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. 2025 Jul 21;247(6):2601–2615. doi: 10.1111/nph.70381

Triphenylphosphonium is an effective targeting moiety for plant mitochondria

Shani Lazary 1, Gal Maman 1, Jenia Binenbaum 1, Mordechai Ronen 1, Iris Tal 1, Elon Yariv 2, Shir Ben Yaakov 1, Eilon Shani 1, Roy Weinstain 1,✉
PMCID: PMC12371181  PMID: 40686066

Summary

  • Small signalling molecules regulate a wide range of physiological and developmental processes in plants, often acting in specific spatial contexts. However, the application of such molecules – whether endogenous or synthetic – typically lacks subcellular resolution, limiting the ability to dissect their localized effects. In this study, we explored the use of triphenylphosphonium (TPP) as a mitochondrial‐targeting motif in plants, using Arabidopsis thaliana and several other species as models.

  • We synthesized and applied fluorescent TPP conjugates and analysed their distribution using confocal microscopy. To evaluate functional specificity, we designed a TPP–ciprofloxacin (CFX) conjugate and compared its activity to that of free CFX in intact plants.

  • Fluorescent TPP conjugates consistently accumulated in mitochondria, bypassing coexisting plastids. The TPP–CFX conjugate inhibited mitochondrial DNA gyrase without affecting the chloroplast isoform, slowed plant growth, elevated mitochondrial reactive oxygen species, and induced nuclear stress‐response genes, whereas free CFX perturbed both organelles.

  • Our results establish TPP as an effective and generalizable tag for mitochondrial targeting in plant systems. This approach enables precise, organelle‐specific chemical manipulation in both model and nonmodel species, offering a new tool for plant cell biology and potential applications in precision agriculture.

Keywords: DNA gyrase, mitochondria, mitochondrial retrograde signalling, organelle targeting, triphenylphosphonium

Introduction

Signalling small molecules are instrumental in shaping plants physiology and adaptive growth (Santner et al., 2009; Brown et al., 2013; Chagas et al., 2018; Waszczak et al., 2018). Most signalling molecules in plants regulate and influence multiple processes, having distinct molecular and phenotypic effects in different cells and even in cellular compartments (e.g. auxin, Bargmann et al., 2013; Savotchenko et al., 2015; gibberellins, Ubeda‐Tomás et al., 2009; Matías‐Hernández et al., 2016; cytokinins, Antoniadi et al., 2015; Immanen et al., 2016), hydrogen peroxide (Rosenwasser et al., 2011; Ozgur et al., 2015), calcium (Kiegle et al., 2000; Kanchiswamy et al., 2014), nitrogen sources (Gifford et al., 2008), lipids (Barbaglia & Hoffmann‐Benning, 2016; Koenig et al., 2020) and pathogenic molecules (Coker et al., 2015). The diversity of functions each of them displays, compounded by the differential responses they invoke in different locations, requires techniques that operate at high resolution to dissect their distinct roles. Yet, in contrast to the remarkable progress made in the manipulation of DNA, RNA and proteins, the ability to manipulate responses to small molecules in plants has remained largely the same; they are invoked by the indiscriminate application of the small molecule to the whole plant, or at the organ level at most, which leads to undirected biological activity that is dependent solely on the molecule's biodistribution pattern. By allowing small molecules to freely distribute within the plant, the spatial context in which they often act is completely lost.

Although synthetic biology has proved useful for the biosynthesis of some endogenous, small organic molecules in planta with spatial and/or temporal control (Barker et al., 2021; Ding et al., 2021), such genome manipulation is often accompanied by unintended developmental consequences and is an effort‐intensive, ad hoc solution. Furthermore, the number of plant species that are amenable for genetic engineering is limited, and the strategy is strictly restricted to those molecules for which a complete suite of biosynthetic enzymes is known and available. The same is true for many valuable synthetic small molecules in plant research, such as specific inhibitors and activity modulators, which are becoming increasingly more available thanks to progress in chemical‐genomics screens (Rigal et al., 2014; Serrano et al., 2015). Such molecules can be applied in a conditional, dose‐dependent and reversible manner, with the advantage of circumventing the limitations of lethality and functional redundancy inherent to mutants (Tóth & van der Hoorn, 2010); yet, their potential utility is far from being fully realized due to the lack of ability to target them to specific locations. The long‐standing inability to target bioactive molecules to specific subcellular locations also has implications in agriculture; only a fraction of an agrochemical applied in the field reaches the site of action in crops (Pimentel & Levitan, 1986; Pimentel, 1995; Wang & Liu, 2007; Mendes et al., 2022) (typically, a specific cellular compartment), resulting in inefficient utilization of resources and significant environmental impact.

In recent years, several efforts have been made to facilitate targeted delivery of small molecules in plants, either by utilizing light‐mediated techniques (Wexler et al., 2019; Hemelíková et al., 2021; Li et al., 2023) or via chemical targeting motifs, to cells and organelles such as the chloroplast (Santana et al., 2020, 2022) (Chl), vacuole (Michels et al., 2020) and cell membranes (Michels et al., 2020), highlighting the feasibility and the potential utility of this approach in both basic and applied plant research. We hypothesized that validating a generic mitochondria targeting motif in plants will engender the development of valuable tools to study this important organelle (Møller et al., 2021) and its components in live, whole plants. Although several mitochondria targeting motifs have been characterized in mammalian cells, none has been comprehensively validated in plants. Out of these known mitochondria targeting motifs (Wang et al., 2021), we chose to focus on triphenylphosphonium (TPP), mainly due to its broad scope of efficacy in organisms other than mammals (Severin et al., 2010; Cochemé Helena et al., 2011; Ng et al., 2014; Wang et al., 2023), its structural simplicity, and its inherent hydrophobicity that could support efficient uptake into plants. The selective accumulation of TPP and other similar lipophilic cations in mitochondria is driven by the plasma membrane potential (Ross Meredith et al., 2008) (ΔΨp = 30–90 mV, negative inside) and the unique mitochondrial inner membrane potential (Ross et al., 2005) (ΔΨm = 120–180 mV), with the negative potential residing on the matrix side (Zielonka et al., 2017). However, unlike most other organisms, plants contain chloroplasts, a photosynthetic double‐membrane organelle that exists in most plant cells in multiple copies. The outer membrane (envelope) potential in chloroplasts is negligible, consistent with its function to allow metabolite exchange between the chloroplast intermembrane space and the cytosol. The inner membrane potential was measured to be 20–50 mV, although under some conditions, a value of 110 mV was obtained (Wu et al., 1991; Szabò & Spetea, 2017), with the negative potential residing on the stroma side. The membrane potential of the thylakoids is reported to be 10–50 mV (Rottenberg & Grunwald, 1972; Joliot & Joliot, 1989), with the negative potential residing, again, on the stroma side. Thus, TPP accumulation in the chloroplast stroma might compete with its accumulation in the mitochondria. Indeed, several studies on isolated chloroplasts have demonstrated the ability of TPP to penetrate into this organelle (Demmig & Gimmler, 1983; Wu et al., 1991).

In this work, we evaluate the distribution pattern of TPP in whole, live plants via fluorescence labelling, confirming its selectivity to the mitochondria in this organism. We then show that TPP can be further leveraged to construct targetable tools to study this important organelle in plants.

Materials and Methods

Plant material and growth conditions

Arabidopsis thaliana (Columbia ecotype) cell culture (Barkla et al., 2014) was kindly provided by Prof. Eilon Shani (Tel Aviv University) and was kindly maintained by Prof. Adi Avni (Tel Aviv University). All A. thaliana lines used in this work are Colombia background (Col‐0 ecotype, Salk Institute, La Jolla, CA, USA). Sterilized seeds were plated on Murashige & Skoog (MS) × 0.5 (Duchefa Biochemic) medium containing 1% w/v sucrose and 0.5% w/v plant agar (Duchefa Biochemic); pH was adjusted to 5.6–5.8 with 1 M KOH. Plates with seeds were stratified for 48 h at 4°C, then transferred to a growth chamber (Percival CU41L5) at 21°C, 100–120 μE m−2 S−1 light intensity under long‐day conditions (16 h light : 8 h dark). For leaf imaging, plants phenotypic characterization and transmission electron microscopy (TEM) experiments, A. thaliana seeds were sown on wet soil, stratified for 48 h at 4°C, then transferred to a growth room at 21°C under long‐day conditions.

Arabidopsis thaliana p35S::H2B‐RFP (Boisnard‐Lorig et al., 2001) transgenic seeds were kindly provided by Prof. Eilon Shani (Tel Aviv University).

Nicotiana benthamiana plants were kindly provided and maintained by Prof. Adi Avni (Tel Aviv University). Seeds were sown on wet soil and transferred to the growth room at 25°C under long‐day conditions.

Lamium amplexicaule seeds were kindly provided by Prof. Nir Ohad (Tel Aviv University). Aegilops longissima and Setaria viridis seeds were kindly provided by Dr. Nir Sade (Tel Aviv University). Lamium amplexicaule, A. longissima and S. viridis seeds were sown on wet Watman Paper placed in a plastic magenta box that was left in the growth room at 25°C under long‐day conditions.

Cloning and expression of GFP in Arabidopsis thaliana's mitochondria

Arabidopsis thaliana p35S:UBP271‐71‐GFP‐BLRP was constructed as follows: p35S:UBP271‐71‐GFP‐BLRP constructed utilizing the Gibson method (NEB‐E2611). The first 71 amino acids of UBP27 (AT4G39370) (Deal & Henikoff, 2010) were amplified from cDNA:

  • F primer: tctagtcgacctgcaggcggccgcaATGGTTTCTAGAAGAGGCTC

  • R primer: ctcacagcggccgcAAAAGAATCGTCTCCGGAG

eGFP‐BLRP amplified from NTF (Pan et al., 2014):

  • F primer: agacgattcttttgcggccgctGTGAGCAAGGGCGAGGAG

  • R primer: ggagaccggcacactggccatatcgTCAAGATCCACCAGTATCCTCATG

Fragments were assembled into pK2GW7 backbone and linearized with SpeI and BstXI.

Seed sterilization

Arabidopsis thaliana seeds from all lines were sterilized by vapour phase sterilization (chlorine fumes). Seeds were placed in Eppendorf tubes inside a sealed desiccator, in the presence of 4% and 32% hydrochloric acid in 11% sodium hypochlorite. After 2–3 h, seeds were taken out and left to ventilate for 30 min in the biological hood. Aegilops longissimum seeds were sterilized by incubation in a 3% sodium hypochlorite solution for 3 min. The sterilized seeds were then washed five times with double distilled water (DDW). Lamium amplexicaule and S. viridis seeds were sterilized by incubation in a 3% sodium hypochlorite solution for 1 min. The sterilized seeds were then washed five times with DDW.

General imaging

All samples were imaged on a laser scanning confocal microscope (Zeiss LSM 780 inverted microscope), with 63×/1.15W or 40×/1.2W objective lenses with transmitted‐light photomultiplier tube (T‐PMT) module for bright field. Hoechst 33342 and 4 were excited with a 405 nm laser; emissions were collected between 410 nm and 556 nm. 1, 3, MitoTracker green and p35s::UBP27‐GFP‐BLRP|BirA transgenic lines were excited with a 488 nm laser; emissions were collected between 493 nm and 556 nm. 2′,7′‐dichlorodihydrofluorescein diacetate (DCF) was excited with a 488 nm laser; emission was collected between 501 nm and 576 nm. 2 and MitoTracker red CMXRos were excited with a 561 nm laser; emissions were collected between 570 nm and 632 nm. p35s:H2B‐RFP transgenic plants were excited with a 561 nm laser; emission was collected between 582 nm and 754 nm. Chlorophyll was excited with a 633 nm laser; emission was collected between 647 nm and 721 nm.

Fluorescent probes imaging

Arabidopsis thaliana

Cell culture was incubated with MitoTracker red (1 μM) and 1 (25 μM) for 30 min, then washed twice with DDW, mounted on a slide and imaged.

Roots: Seedlings were grown on MS agar. At 5–7 d, seedlings were incubated in liquid MS with the inspected probes: MitoTracker red (1 μM) and 1 (25 μM); MitoTracker red (1 μM) and 3 (10 μM); MitoTracker green (10 μM) and 2 (10 μM); MitoTracker red (1 μM) and fluorescein (25 μM); MitoTracker red (1 μM) and 7 (10 μM) and MitoTracker green (10 μM) and TAMRA (10 μM). The seedlings were left for 3 h then washed twice with DDW and taken for imaging.

For all probes, colocalization analysis was performed using the Coloc2 function in Fiji software. Noncorrected and nonthresholded images of probes and respective MitoTracker were loaded to the software. Pearson correlation coefficients and Manders correlation coefficients above the autothreshold (tM1 and tM2) were determined from ROI (n ≥ 5).

p35S::UBP27‐GFP‐BLRP transgenic A. thaliana seedlings were grown on MS agar. At 5–7 d, seedlings were incubated for 3 h in liquid MS with 2 (10 μM), then washed twice with DDW and taken for imaging. A line was drawn over an individual mitochondrion and grey values were measured using Fiji.

Leaves: using compound 1—Seedlings were placed in 3 ml wells containing 1 (25 μM) in liquid MS and were vacuum infiltrated for 10 min, then MitoTracker red (1 μM) was added and the seedlings left for incubation. After 3 h, seedlings were washed twice with DDW and taken for imaging.

Using compounds 2 and 3—Seedlings were grown on MS agar plates. At 5–7 d, seedlings were incubated in liquid MS with the inspected probes: MitoTracker red (1 μM) and 3 (10 μM) or MitoTracker green (10 μM) and 2 (10 μM). The seedlings were left for 3 h, then washed twice with DDW and taken for imaging.

Lamium amplexicaule, Aegilops longissimum and Setaria viridis

Seedlings were grown in magenta boxes on MS agar medium. At 5–7 d, seedlings were incubated for 3 h in liquid MS with MitoTracker red (1 μM) and 3 (10 μM), then washed twice with DDW and taken for imaging.

Cross sections

Arabidopsis thaliana cross sections were generated as described before (Skopelitis et al., 2018). Briefly, a 3‐wk‐old Arabidopsis plant was incubated for 12 h in 0.5% liquid MS containing MitoTracker green (10 μM) and 2 (10 μM). The plant hypocotyl was fixed in 4% paraformaldehyde dissolved in 1× phosphate buffered saline (PBS) supplemented for 60 min. Fixed tissues were washed twice (10 min per wash) in 1× PBS and were then transferred to ClearSee solution. After 12 h, the tissues were embedded in 8% Low Melting Agarose (Invitrogen) for 15 min and were then sectioned using a VT1000S vibratome (Leica) to generate 100 μm sections. Sections were then taken for imaging.

Carbonyl cyanide‐p‐trifluoromethoxyphenylhydrazone uncoupling

Arabidopsis thaliana seedlings were grown on MS agar. At 5–7 d, seedlings were incubated for 3 h in liquid MS with 3 (10 μM) and MitoTracker red (1 μM). For the last hour of the incubation, FCCP (10 μM) was added. At the end of the incubation, seedlings were washed twice with DDW and subsequently imaged.

Emission of 4 in the absence or presence of DNA

Total DNA, extracted from A. thaliana, was added incrementally to a cuvette containing 4 (5 μM) in DDW. Fluorescence emission spectrum (λ ex = 348 nm, λ em = 358–650 nm) was recorded after each addition. Data were processed using OriginPro software.

Imaging of 4 in Arabidopsis thaliana roots

p35S:H2B‐RFP or p35S::UBP27‐GFP‐BLRP transgenic A. thaliana seedlings were grown as described before. At 5–7 d, seedlings were incubated for 3 h in liquid MS with Hoechst 33342 (2.5 μM) or 4 (5 μM), washed twice with DDW and subsequently imaged.

Modelling of ciprofloxacin analogues in complex with GyrA/GyrB

The structures of GyrA and GyrB from A. thaliana (Uniprot ID: Q9CAF6 and Q9SS38) were modelled with Alphafold 2.0 [PMID: 34265844], using the X‐ray crystallography structure of the same proteins from Staphylococcus aureus (PDB ID: 2XCS, PMID: 20686482). The resulting computational model was then superimposed with the experimental model, thereby aligning the computational model with the dsDNA helix, which constitutes a large portion of the CFX‐binding site.

The computational model, along with the dsDNA, was then prepared using Protein Preparation Wizard (Schrodinger LLC, New York, NY, USA) [PMID: 23579614]. CFX, 5, and 6 were prepared for docking using ligprep [PMID: 17899391]. Docking of these molecules was done with Glide Standard Precision mode [PMID: 15027865, 15027866], in two stages. Initially, CFX was docked with no constraints into the binding site. The highest scoring pose had CFX sandwiched between the nitrogen bases of the dsDNA. This pose was used as a core constraint, with a maximum of 0.10 Å root‐mean‐square deviation (RMSD) tolerated between the core and the matching sections of 5 and 6. This was necessary because of the long flexible tail added to the analogues, which makes exhaustively sampling all the different conformations computationally prohibitive, and the unconstrained docking results were therefore poor in quality.

Gyrase inhibition assay

Gyrase inhibition assay was performed using TopoGEN Escherichia coli DNA gyrase and relaxed DNA assay kit (TG2000G‐1KIT) according to the manufacturer's instructions.

Plants phenotypic characterization

Arabidopsis thaliana seeds were sown and grown in pots. On day 10 and day 12, each pot was sprayed with 1 ml DDW containing 0.05% TWEEN 20 (Merk) and CFX (10 μM) or 6 (50 μM) or 3 (50 μM). Morphological and photosynthesis parameters were analysed with the PlantScreen™ Phenotyping System, Photon Systems Instruments (PSI), Czech Republic. Plants were sown in PSI standard pots and imaged once a day from day 14 to day 26. On the last measuring day, plants were also photographed using a Nikon D5300 with an AF‐S micro Nikkor 60 mm lens.

Root elongation assay

Arabidopsis thaliana seedlings were grown on MS agar as described in ‘Plant material and growth conditions’ in the Materials and Methods section. On day 5, the seedlings were transferred to MS plates containing various concentrations of CFX, 6 or 3. The seedlings were placed in a growth chamber under long day condition and root length was marked daily. On day 10, the plants were scanned, and root length was measured using Fiji software.

Transmission electron microscopy imaging

Arabidopsis thaliana plants were grown in pots. On day 10 and day 12, each pot was sprayed with 1 ml DDW containing 0.05% TWEEN 20 (Merk) and CFX (10 μM) or 6 (50 μM) or 3 (50 μM). On the day 14, c. 1 × 1 cm2 tissues were cut from leaves. Tissues were fixed in 2.5% glutaraldehyde in PBS overnight at 4°C. After several washings in PBS, tissues were postfixed in 1% OsO4 in PBS for 2 h at 4°C. Dehydration was carried out in graded ethanol followed by embedding in glycid ether. Thin sections were mounted on Formvar/carbon‐coated grids, stained with uranyl acetate and lead citrate and examined in Jeol 1400 – Plus transmission electron microscope (Jeol, Japan). Images were captured using SIS Megaview III and iTEM the Tem imaging platform (Olympus).

Relative DNA copy number

Arabidopsis thaliana seedlings were grown on MS agar plates. On the day 5, seedlings were transferred onto MS plates containing CFX (10 μM) or 6 (50 μM) for 24 h. In this study, c. 100 seedlings were harvested, frozen in liquid nitrogen and crushed by a TissueLyser to form a thin powder. Five hundred microlitres phenol–chloroform (1 : 1) buffer were added to each sample, mixed well and centrifuged for 3 min. The supernatant was transferred to a new tube with an equal volume of chloroform to remove phenol traces, centrifuged for 3 min, and the supernatant was transferred to a new tube. An equal volume of 5 M ammonium acetate was added to the supernatant to a final concentration of 2.5 M, followed by the addition of 2–2.5 ml of cold 100% EtOH. Then, samples were stored at −20°C for 12 h and centrifuged at 16 900  g for 15 min. Supernatants were decanted. One millilitre of 70% EtOH was added to the pellet, and the mixture was centrifuged for 1 min. The supernatant was discarded, and the pellets were air‐dried for 10 min and resuspended in TE buffer. Quantitative reverse transcription polymerase chain reaction was performed with 10 ng DNA in a final volume of 10 μl with Fast SYBR™ Green Master Mix (Applied Biosystems, Cat. No. 4385612) using StepOnePlus™ System and software (Thermo Fisher Scientific). The reaction conditions included 40 amplification cycles (3 s at 95°C, 30 s at 60°C). Three technical repeats were performed for each DNA sample, and at least three biological repeats were used for each treatment. The relative quantification was calculated with the ΔΔCt method; TUB2 (TUBULIN BETA CHAIN 2) and ACT2 (ACTIN 2) were used as reference genes. Primers are specified in Supporting Information Table S1.

2′,7′‐dichlorodihydrofluorescein diacetate staining

Nicotiana benthamiana plants were grown in pots. At c. 2 wk old, leaves were infiltrated with 6 (50 μM) or DDW. After 3 h, leaves were infiltrated again with MitoTracker red (1 μM) and DCF (50 μM). After 10 min, leaves were imaged. In each image, five mitochondria and five chloroplasts were chosen randomly, and DCF emission was measured using zen 3.1 blue edition software.

Relative RNA expression

Arabidopsis thaliana seedlings were grown on MS agar plates. On day 5, seedlings were transferred onto MS plates containing CFX (10 μM) or 6 (50 μM) or mock treatment. After 3 or 6 h, seedlings were harvested and grounded in liquid nitrogen. Total RNA was extracted from 70 to 100 mg grounded plant tissue using RNeasy Plant Mini Kit (QIAGEN, Cat. No. 74904), according to the manufacturer protocol. DNA was removed by DNase I, RNase‐free (Thermo Fisher Scientific, Cat. No. EN0521). Total RNA (2 μg) was converted to complementary DNA (cDNA) using High‐Capacity cDNA Reverse Transcription Kit (applied biosystems by Thermo Fisher Scientific, Cat. No. 4368814) according to manufacturer protocols. Quantitative PCR was performed with 40 ng cDNA in a final volume of 20 μl, with Fast SYBR™ Green Master Mix (Applied Biosystems, Cat. No. 4385612) using the QuantStudio1 Real‐Time PCR System (Thermo Fisher Scientific). The reaction conditions included 40 amplification cycles (3 s at 95°C, 30 s at 60°C). Three technical repeats were performed for each cDNA sample, and three biological repeats were used for each treatment. Relative quantification was calculated using the ΔΔCt method, with PP2AA3 (PROTEIN PHOSPHATASE 2A SUBUNIT A3) used as the reference gene. Primers are specified in Table S2.

Data analysis

All graphs (line charts, bar charts, box plots) were generated using OriginPro 2024. Statistical analyses were performed either with Microsoft Excel (t‐test) or OriginPro2024 (ANOVA). Colocalization analyses were done using Fiji. Grey value measures were done either with Fiji (fluorescent probes imaging) or ZEN 3.1 blue edition (DCF staining). EC50 calculations (DoseResp fitting curve) were performed using OriginPro2024. Figures were assembled using Photoshop.

Synthetic methods and characterization

Synthesis and characterization of all reported compounds are included in Supporting Information Methods S1.

Results

Triphenylphosphonium selectively accumulates in plants mitochondria

The cellular distribution pattern of TPP was examined using fluorescently labelled conjugates, synthesized in three steps from TPP (Fig. 1a, compounds 1–3). The labelling fluorophores differ mostly in their electronic charge, intending to evaluate the effect of the cargo properties on parameters such as tissue penetration and distribution of the conjugate.

Fig. 1.

TPP probes accumulate only in Arabidopsis mitochondria, verified by confocal co-labelling and co-localization plots.

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Fluorescent conjugates of triphenylphosphonium (TPP) accumulate in mitochondria of Arabidopsis thaliana cell cultures and seedlings. (a) Structures of synthesized TPP conjugates and (b) representative confocal images of A. thaliana cultured cells after incubation with 1 (25 μM, 30 min, green) and MitoTracker red (1 μM, 30 min, magenta). (c) Box plot chart presenting colocalization between mitochondria (Mit) and chloroplasts (Chl) in cells using Pearson correlation coefficients (PCC) of nonthresholded images as shown in (b). (d) Representative confocal images of 5‐d‐old A. thaliana seedlings roots incubated for 3 h with MitoTracker red (1 μM, magenta) or green (10 μM, green) and the corresponding fluorescent probes 1 (25 μM, green) or 2 (10 μM, magenta) or 3 (10 μM, green). (e) Representative confocal images of 5‐d‐old A. thaliana leaves. For 1 (green), seedlings were vacuum infiltrated for 10 min, then left for 3 h incubation with MitoTracker red. For 2 (magenta) and 3 (green), seedlings were incubated for 3 h with MitoTracker red (1 μM, magenta) or green (10 μM, green) and the corresponding fluorescent probes 2 (10 μM, magenta) or 3 (10 μM, green). (f) Box plot chart presenting colocalization analysis in roots for each probe and its co‐responding MitoTracker using PCC of nonthresholded images as shown in (d). (g) Box plot chart presenting colocalization analysis in leaves for each probe and its co‐responding MitoTracker and Chl natural fluorescence, using PCC of nonthresholded images as shown in (e). Bars: (b, d and e), 5 μm. (c, f and g) Horizontal lines represent the means and box limits indicate the 25th and 75th percentiles. Whiskers indicate ± SD.

In light‐cultured A. thaliana cells, conjugate 1 accumulated selectively in punctate structures that colocalized with the mitochondria marker MitoTracker red and was absent from any other organelle, including plastids (Fig. 1b,c). Similar results were observed when wild‐type (WT) A. thaliana seedlings (Col‐0) were incubated in solutions of 1–3; the fluorescently labelled TPP derivatives accumulated exclusively in punctate structures that were colabelled by either a green or a red MitoTracker marker in both root and leaf cells (Fig. 1d–g). In comparison, the respective free dyes showed mostly nonspecific cellular distribution in these tissues (Fig. S1). The identity of the punctate structures in which the fluorescently labelled TPP accumulated as mitochondria was further solidified by their colocalization with a GFP fusion of the mitochondria membrane protein UBIQUITIN‐SPECIFIC PROTEASE 27 (UBP27) in transgenic Arabidopsis seedlings (p35S::UBP27‐GFP‐BLRP, Fig. 2a). Notably, the fluorescent signal of 2 was observed in the organelle interior (Fig. 2b), suggesting it accumulates in the mitochondria matrix in accord with the large negative membrane potential across the inner mitochondrial membrane. In accordance, cotreatment of WT seedlings with 3, MitoTracker red and the uncoupler carbonyl cyanide‐p‐trifluoromethoxyphenylhydrazone (FCCP) led to complete loss of the punctate structures labelling observed in the absence of FCCP (Fig. S2), suggesting that the uptake and retention of the TPP conjugates in the mitochondria are dependent on intact mitochondria membrane potential.

Fig. 2.

TPP conjugate fluorescence marks Arabidopsis mitochondria and matrix, including in hypocotyl cells.

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Fluorescent conjugates of triphenylphosphonium (TPP) accumulate in Arabidopsis thaliana mitochondria matrix. (a) Representative confocal images of 5‐d‐old p35S::UBP27‐GFP‐BLRP transgenic A. thaliana seedlings incubated with 2 (10 μM, 3 h, magenta). (b) Line chart representing the grey value in the mitochondrion labelled in A, showing accumulation of 2 in a mitochondrion interior. (c) Representative confocal images of a 1‐month old A. thaliana plants hypocotyl cross section. Only the roots of the plants were treated for 16 h with 2 (10 μM, magenta) and MitoTracker green (10 μM, green), then hypocotyls were fixed in agarose, sectioned and imaged. Orange box shows borders of an area shown as an enlarged image on the right in a and c.

The effect of the cargo charge on TPP uptake and distribution pattern was studied before in mammalian cell cultures but is distinct in plants; TPP conjugates with a negative‐ or a neutral‐charge cargo (1 and 3, respectively) were highly specific to the mitochondria. However, c. 2.5‐fold higher concentration of 1, bearing a negatively charged cargo, was required to obtain a detectable signal in planta compared to the neutral‐cargo bearing 3, suggesting a lower uptake of the former. In comparison, zwitterionic TPP conjugates showed higher cellular and mitochondrial uptake in mammalian cells (Finichiu et al., 2013; Pala et al., 2020). A cargo with a positive charge (2) led to some association of the conjugate with the cell wall (Figs 1d, 2a), presumably with the pectin‐rich primary cell wall (Caffall & Mohnen, 2009). Interaction of positively charged species with the cell wall is well documented (MacDougall et al., 2001; Rounds et al., 2011; Mravec et al., 2014). While these cellular structures (i.e. cell wall and mitochondria) can be easily differentiated visually, the effect should be considered when the cargo is nonfluorescent.

To assess the mobility of TPP conjugates in planta and their ability to reach deep tissues, only the roots of 1‐month‐old Arabidopsis plants were dipped in a solution of 2 (10 μM) and MitoTracker green (10 μM) for 16 h, followed by fixation and agarose embedment. Cross sections of the hypocotyl revealed that compound 2 translocated acropetally and accumulated in mitochondria of inner‐layer cells (Fig. 2c).

To evaluate the biodistribution of TPP conjugates in other plant species that are not readily amenable to genetic manipulations, compound 3 (10 μM) was applied to seedlings of L. amplexicaule (henbit), A. longissimi (a wheat relative) and S. viridis (green foxtail) in solution, and their hypocotyls (which contain chloroplasts) were consequently imaged. The results recapitulated those observed in Arabidopsis: selective accumulation of 3 in punctate structures that colocalized with MitoTracker red (Fig. S3). A. longissimi characteristics allowed for more comprehensive imaging, showing accumulation in different plant organs, such as root hairs and the hypocotyl (Fig. S4).

Collectively, these data establish that TPP conjugates are readily taken up by plants, translocate and accumulate in the mitochondria, including in photosynthetic organs, and suggest they can be utilized to localize molecules of interest selectively in the mitochondria of live plants. We therefore next evaluated the potential utility of TPP in plants for the development of mitochondria‐selective effectors.

A triphenylphosphonium‐Hoechst dye conjugate selectively labels mtDNA in planta

Mitochondrial DNA (mtDNA) plays important roles in diverse physiological processes and its direct imaging is an indispensable molecular tool for their studying (Prole et al., 2020). Several DNA‐intercalating dyes are used for imaging of mtDNA by diffraction‐limited microscopy, such as 4′,6‐diamidino‐2‐phenylindole (DAPI) (Dellinger & Gèze, 2001; Shen et al., 2019), SYBR dyes (Arimura et al., 2004; Jevtic et al., 2018) and picoGreen (Ashley et al., 2005; He et al., 2007), including in plants (Arimura et al., 2004). However, these dyes have a substantial drawback as they stain all DNAs of the cell, and therefore require an elaborate treatment or image analysis to avoid the significant fluorescence emanating from the abundant nuclear DNA. We speculated that conjugation of a fluorogenic DNA intercalator with a mitochondria‐trappable entity might afford a ‘turn‐on’ fluorescent stain suitable for specific mtDNA imaging.

To test this concept, we synthesized a TPP conjugate that resembles the Hoechst 33342 dye (compound 4, Fig. 3a), one of the most prevalent DNA markers. The strong affinity of this dye family to DNA (binding constant of c. 108 M−1) (Wiederholt et al., 1997) has been previously leveraged for targeted delivery of conjugates to mammalian cells nuclei (Lukinavičius et al., 2015; Bucevičius et al., 2018; Hara et al., 2018; Makanai et al., 2024). Conjugate 4 exhibited a DNA‐dependent fluorescence emission enhancement (Fig. 3b), typical of the Hoechst dyes family (Cosa et al., 2001), validating that the conjugation to TPP does not interfere with the dye's ability to intercalate within DNA. This observation is in agreement with a prior report on the ability of an oxidized ethidium–TPP conjugate (MitoSOX) to bind DNA (Shchepinova et al., 2017). As expected, application of the free Hoechst dye 33342 to transgenic Arabidopsis seedlings expressing H2B‐RFP (35S::H2B‐RFP) as a fluorescent nuclear marker resulted in exclusive staining of nuDNA, as evident by the colocalization of the two fluorophores (Fig. 3c). In comparison, following the application of 4 to similar seedlings, the fluorescence emission was observed exclusively in intracellular punctate structures and was completely devoid from the nucleus. The identity of these punctate structures as mitochondria was independently validated in transgenic seedlings expressing a genetically encoded mitochondria marker (p35S::UBP27‐GFP‐BLRP) (Fig. 3d).

Fig. 3.

TPP-Hoechst dye highlights mitochondrial DNA but not nuclei in live Arabidopsis seedlings.

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A triphenylphosphonium (TPP)‐Hoechst dye conjugate selectively stains mtDNA in planta. (a) Structure of synthesized Hoechst dye 33 342 conjugate with TPP 4. (b) Fluorescence emission of 4 with increasing amount of DNA following excitation at 348 nm. (c) Representative confocal images of 5‐d‐old p35S:H2B‐RFP (magenta) transgenic Arabidopsis thaliana seedlings incubated with Hoechst 33342 (2.5 μM, 3 h, blue) or 4 (5 μM, 3 h, blue). (d) Representative confocal images of 5 d p35S::UBP27‐GFP‐BLRP (green) transgenic Arabidopsis thaliana seedlings incubated with Hoechst 33342 (2.5 μM, 3 h, blue) or 4 (5 μM, 3 h, blue). The orange box in c and d indicates the region shown at higher magnification in the image on the right.

These results not only reinforce the specificity of TPP accumulation in plants mitochondria but also establish the ability of the conjugated cargo to access the mitochondria matrix and to further interact with endogenous entities within it.

A triphenylphosphonium–ciprofloxacin conjugate selectively inhibits mitochondrial gyrase

To take further advantage of this capability, we sought to deliver an enzyme inhibitor specifically to the mitochondria matrix. Small molecule inhibitors are an indispensable tool for dissecting cellular functions. Nevertheless, they typically lack the spatial specificity that would allow them to differentiate between targets that are present in multiple cellular organelles. For example, DNA gyrase is a type‐II topoisomerase that has key roles in DNA replication and transcription (Vos et al., 2011) and that localizes in plants to both the chloroplast and the mitochondria (Wall et al., 2004). The specific functions of this enzyme in plants are not clearly understood, as most eukaryotes do not express it. Previous studies have shown that knockout of the gyrase genes leads to embryo‐ or seedling‐lethal phenotypes, impeding analysis of their function through traditional genetic tools (Wall et al., 2004). Quinolone and aminocoumarin antibiotics were shown to be effective inhibitors of plant gyrases (Wall et al., 2004; Evans‐Roberts et al., 2016) and are even being explored as potential novel herbicides (Wallace et al., 2018). However, both inhibitors lack the spatial specificity that would allow dissecting the differential roles of DNA gyrase in the chloroplasts and in the mitochondria. We hypothesized that a mitochondria‐targeted ciprofloxacin (CFX), a known gyrase inhibitor of the quinolone family (Hooper, 2001), would distinguish between mitochondrial and chloroplast gyrases and help gain a better understanding of its mitochondria‐specific roles.

To this end, we synthesized two derivatives of CFX–TPP conjugates, differing in the functional group connecting the two units (5 and 6, Fig. 4a). Both alkylation (5) and acylation (6) of the piperazine secondary amine were previously utilized for derivatization of CFX and were shown to not significantly interfere with its mechanism of inhibition (Zhang et al., 2018). Computational docking experiments showed no significant differences in the estimated binding affinities and that the binding modes were conserved for the CFX part (Fig. 4b–d). The aliphatic addition lies near the DNA coil and does not cause allosteric clashes. These results were observed for both the bacterial gyrase (Staphylococcus aureus, 2XCS crystal structure) and for the two Arabidopsis gyrase complexes (AtGyrA – At3g10690 and chloroplast AtGyrB – At3g10270 or mitochondrial AtGyrB – At5g04130, using alpha‐fold2 structural predictions based on the 2XCS experimental data), which show high similarity in their sequence to the bacterial enzyme and that were shown to bind CFX (Evans‐Roberts et al., 2016). Both compounds showed c. 30% retention of gyrase inhibition activity compared to free CFX against the E. coli gyrase in a relaxation of supercoiled DNA assay in vitro, with compound 6 showing higher potency at higher concentrations (Fig. 4e). In comparison, conjugate 3 did not have any measurable effect on gyrase activity in this assay, suggesting that the gyrase inhibition observed with compounds 5 and 6 is CFX‐dependent (Fig. S5). Based on these results, we focused further efforts on the more potent compound 6.

Fig. 4.

CFX-TPP analogues dock to plant gyrase and retain ~30% inhibitory activity in vitro.

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Triphenylphosphonium (TPP) conjugates of ciprofloxacin (CFX) retain inhibitory activity against DNA gyrase. (a) Structures of synthesized CFX–TPP conjugates. (b–d) Three‐dimensional representations of CFX (green), CFX–TPP conjugates 5 (magenta) and 6 (blue) molecular docking poses in (b) Staphylococcus aureus, gyrase crystal structure (PDB ID: 2XCS) or alpha‐fold2 models of Arabidopsis thaliana (c), chloroplasts gyrase (Gyr) and (d) mitochondrial gyrase (e), gyrase inhibition assay and bar chart of gyrase relative inhibition. Whiskers indicate ± SD. Significance was determined by one‐way ANOVA analysis with Tukey's honestly significant difference (HSD) post‐hoc test. Different capital letters (A–E) indicates a significant difference at a P value of < 0.05.

A root elongation assay performed on 5‐d‐old WT Arabidopsis (Col‐0) seedlings showed that 6 has an EC50 value of c. 10 times higher than CFX (16.2 vs 1.4 μM, respectively, Figs 5a, S6A,B), in agreement with the results obtained for inhibition of the E. coli gyrase. Compound 3, which is not expected to have any biological effect, showed an EC50 > 50 μM in this assay (Fig. S7A,B), suggesting that in prolonged exposure to high concentrations, the TPP moiety can exert a phytotoxic effect. To assess the specificity of 6 towards the mitochondrial gyrase in planta, 14‐d old Arabidopsis plants were sprayed with free CFX or 6 (10 or 50 μM, respectively) and subsequently monitored for 14 d. Within the period of observation, plants treated with CFX maintained a similar growth rate to that of untreated ones (Fig. 5b) but showed an abnormal leaves shape and progressive decolouration (Fig. S6C–E), indicative of chloroplast dysfunction. Correspondingly, measured photosynthetic parameters, such as photosystem II quantum yield, showed a significant drop in comparison to nontreated plants (Figs 5C, S6F–H). By contrast, plants treated with 6 maintained photosynthetic parameters similar to those of untreated plants but were significantly smaller, presumably indicative of mitochondria dysfunction below a minimal threshold required to support growth (Hauben et al., 2009). The fact that the phenotypic manifestation of freex CFX effect is observed mostly via chloroplast functions could be a result of its distribution being skewed towards this organelle or that chloroplasts are more sensitive to its effect than the mitochondria. In agreement with the above results, TEM images of plants treated with CFX revealed structurally deformed chloroplasts with altered thylakoid arrangements, as previously reported (Evans‐Roberts et al., 2016) and disrupted mitochondria morphology, whereas 6 treated plants showed normally looking chloroplasts and structurally deformed mitochondria (Fig. 5D). Importantly, WT seedlings and plants treated with the nonbioactive compound 3 did not show any of the phenotypes described above (Fig. S7C–J).

Fig. 5.

Mitochondria-targeted CFX-TPP slows growth and specifically impairs mitochondrial gyrase and ultrastructure.

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A ciprofloxacin (CFX)–Triphenylphosphonium (TPP) conjugate inhibits Arabidopsis thaliana mitochondrial DNA gyrase in planta. (a) Root elongation essay of 5‐d old A. thaliana seedlings grown for 5 d on indicated concentrations of CFX (green) or 6 (blue). Each data point represents an average of n ≥ 8 plants. (b, c) Measurement over 2 wk of A. thaliana plants rosette leaves area (b) and maximal PSII quantum yield (c), after treated twice (on days 10 and 12) with CFX (10 μM, green) or 6 (50 μM, blue) by spray. Each data point represents the mean of 20 plants. (d) Transmission electron microscopy (TEM) images of organelles from 2‐wk‐old A. thaliana plants treated with CFX (10 μM) or 6 (50 μM) for 24 h. Bars, 200 nm. (e) Relative DNA copy number, quantified by quantitative polymerase chain reaction, of 5‐d‐old A. thaliana seedlings treated for 24 h with CFX (10 μM) or 6 (50 μM). Significant was determined by one‐way ANOVA analysis with Tukey's honestly significant difference (HSD) post‐hoc test. Different capital letters (A–C) indicate a significant difference at a P value of < 0.05. For (A, B, C and E), Whiskers indicate ± SE. Chl, chloroplasts; Mit, mitochondria.

A previous study has found that following treatment with CFX, there is a reduction in the number of both chloroplasts and mitochondria in WT plants (Evans‐Roberts et al., 2016). To evaluate the effect of the targeted CFX, 5‐d‐old Arabidopsis plants were treated with either CFX or 6 (10 or 50 μM, respectively) and the amount of mitochondrial and chloroplast DNA (as a proxy for organelles amounts) was quantified 24 h later with respect to the amount of nuclear DNA using quantitative polymerase chain reaction (qPCR) for two genes in each organelle (TUB2 and ACT2, PSBK and YCF3 and COX2 and RPS3, for nuclear, chloroplast or mitochondrial DNA, respectively) (Fig. 5E). Treatment with CFX led to a significant reduction in the amount of both mitochondrial and chloroplast DNA, while treatment with 6 led to a significant reduction in mitochondrial DNA and a nonsignificant (c. 20%) reduction in the amount of chloroplast DNA. These results can suggest that some mitochondria‐targeted CFX ends up in the chloroplast, directly affecting this organelle, or, alternatively, that the reduction observed in the amount of chloroplasts is a downstream effect of the reduction in the amount of functional mitochondria (Busi et al., 2011). Collectively, these results suggest that compound 6 effectively inhibits DNA gyrase specifically in plant mitochondria.

Many stress conditions in plants lead to elevated reactive oxygen species (ROS) levels, which in turn activate various stress‐response pathways (Waszczak et al., 2018). We hypothesized that interfering with mitochondrial replication via gyrase inhibition should lead to some form of a stress response. Indeed, when 6 was injected into N. benthamiana leaves, a significant increase in ROS level, as measured by 2′,7′‐dichlorodihydrofluorescein diacetate (DCF) fluorescence intensity, was observed 3 h post‐treatment in the mitochondria but not in chloroplasts, compared to leaves injected only with water (Fig. 6a,b). This suggests that a mitochondrial mechanism (Waszczak et al., 2018) senses the defunct activity of the gyrase, or its downstream effects, and responds by increasing ROS levels. Moreover, a significant increase in the transcription of ARABIDOPSIS NAC DOMAIN PROTEIN 13 (ANAC013) transcription factor and the downstream ALTERNATIVE OXIDASE 1A (AOX1a) gene was observed 3 h posttreatment with 6 (Fig. 6c). Both genes are common markers of mitochondrial stress responses that are mediated by ROS signalling (Selinski et al., 2024).

Fig. 6.

Mitochondrial gyrase inhibition raises mitochondrial ROS and up-regulates nuclear stress genes.

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Inhibition of mitochondria DNA gyrase leads to nuclear stress response. (a) Representative confocal images of 2‐wk‐old Nicotiana benthamiana leaves treated with 6 (50 μM) or double distilled water (DDW) and incubated with 2′,7′‐dichlorodihydrofluorescein diacetate (DCF) (50 μM, green) and MitoTracker (1 μM, magenta). (b) Box plot presenting the intensity of DCF. Each point represents mean fluorescence intensity of five organelles randomly selected. (c) Box plot presenting the relative RNA expression levels of ANAC013 and AOX1a transcripts, quantified by quantitative reverse transcription polymerase chain reaction, of 5‐d‐old Arabidopsis seedlings treated for 3 h or 6 h with 6 (50 μM). For b and c, centre lines represent the means and box limits indicate the 25th and 75th percentiles. Whiskers indicate ± SD. Significance was determined by one‐tailed Welch t‐test, *, a significant difference at a P value of <0.05, **, a significant difference at a P value of < 0.01. Chl, chloroplasts; Mit, mitochondria.

Discussion

The key outcome of this study is the validation of TPP as a reliable mitochondrial targeting motif in plants. This finding is particularly significant given the unique challenges posed by plant cells, which contain multiple organelles with overlapping structural properties, such as mitochondria and chloroplasts, and various barriers, such as the cell wall, that are not present in mammals. The observed selectivity suggests that TPP‐based strategies could overcome the limitations of traditional small‐molecule applications, which often lack spatial specificity and lead to off‐target effects. Other mitochondria‐targeting motifs are known (Wang et al., 2021), but await further validation in plants.

The study highlights the potential of TPP conjugates for the development of mitochondria‐selective probes and effectors. The successful conjugation of TPP to a Hoechst dye, resulting in specific labelling of mtDNA in planta, should provide a means to study mitochondrial processes without signal interference from nuclear DNA, which is a common challenge when using traditional DNA‐binding dyes. The ability to visualize mtDNA in live, whole plants with minimal background interference will enable more detailed investigations into its roles in plant development and stress responses (Møller et al., 2021). Moreover, to the best of our knowledge, this is the first specific mtDNA‐visualizing agent reported in any organism and is therefore a valuable addition to the growing toolkit of mitochondria imaging probes (Samanta et al., 2019).

The TPP–CFX conjugate, which selectively inhibited mitochondrial DNA gyrase, represents an advancement in the ability to dissect organelle‐specific functions. A similar approach could be applied to study other enzymes that reside in the mitochondria matrix but also in other organelles, such as superoxide dismutase (Shams et al., 2024), aconitase (Hooks Mark et al., 2014) and glutamine synthase (Taira et al., 2004). The differential effects observed between free CFX and the TPP‐conjugated version underscore the importance of spatial specificity in small‐molecule applications. By restricting the activity of CFX to mitochondria, we unveiled phenotypic and molecular effects that arise specifically from inhibiting DNA gyrase in this organelle, mostly avoiding confounding effects from gyrase chloroplast malfunction. The slower growth phenotype observed in seedlings treated with mitochondria‐targeted CFX is in accord with mitochondria dysfunction (Wallström et al., 2014; Moullan et al., 2015; Ayabe et al., 2023). Reproductive tissues are typically the ones most severely impaired by mitochondrial dysfunction, but vegetative development is also often affected, manifesting in slow growth and overall stunted stature (Liberatore et al., 2016). Mutants in other DNA replication machinery components, such as the mitochondrial polymerase IB, show a similar phenotype (Parent et al., 2011). Nevertheless, knockout of the mitochondrial GyrB2 unit results in much more deleterious effects (Evans‐Roberts et al., 2016). This might suggest that mitochondrial gyrase activity is more vital in the first phases of germination and seedling development than in later ones or simply reinforce the importance of a properly functioning mitochondria in these stages (Liberatore et al., 2016). Inhibition of the mitochondrial DNA gyrase was found to elevate ROS levels in this organelle and to upregulate the transcription of ANAC013, a known mediator of mitochondria retrograde regulation (MRR) (De Clercq et al., 2013), as well as AOX1a, by far the most commonly used indicator of mitochondrial retrograde response (Clifton et al., 2006; Rhoads & Subbaiah, 2007). This suggests that improper function of the mitochondrial DNA gyrase is signalled to the nucleus through the endoplasmic reticulum, presumably via ROS as an MRR‐triggering signalling molecule (Vanlerberghe et al., 2002; Rhoads et al., 2006; Khan et al., 2024).

In conclusion, this study demonstrates a novel approach for controlling the bioactivity of small molecules in plants with high spatial resolution by utilizing a targeting chemical moiety. It establishes TPP as a versatile mitochondrial targeting motif in plants, with potential for developing organelle‐specific probes and effectors. In addition, dozens of TPP‐based effectors and probes are used in mammalian systems, and this work opens the way for their implementation in plant research. The successful application of TPP‐based conjugates in various plant species, especially ones that are not amenable to genetic engineering, suggests that this approach could be broadly applicable, offering new avenues for both basic research and agricultural innovation. Finally, the potential for cross‐application of this approach to other organelles (Lin et al., 2021) should allow for an expanded toolkit to dissect and manipulate the functions of distinct organelles.

Competing interests

None declared.

Author contributions

RW and SL conceived the project, and RW supervised the project. SL designed and performed experiments and analysed the data. GM and MR performed qPCR experiments, and MR also designed and performed phenotypic experiments. JB performed cross‐section samples, SBY performed gyrase inhibition assays, and IT constructed the p35S::UBP27‐GFP‐BLRP transgenic lines under the supervision of ES. EY performed computational predictions and docking experiments.

Disclaimer

The New Phytologist Foundation remains neutral with regard to jurisdictional claims in maps and in any institutional affiliations.

Supporting information

Fig. S1 Nontargeted fluorophores do not specifically accumulate in mitochondria.

Fig. S2 FCCP disrupts mitochondrial accumulation of 3.

Fig. S3 Fluorescently labelled TPP localizes to mitochondria in various plant species.

Fig. S4 Fluorescently labelled TPP localizes to mitochondria in multiple organs of Aegilops longissimi.

Fig. S5 TPP moiety does not inhibit bacterial DNA gyrase supercoiling activity in vitro.

Fig. S6 CFX–TPP conjugate inhibits Arabidopsis mitochondrial DNA gyrase in planta.

Fig. S7 TPP moiety does not show phenotypic effects in Arabidopsis thaliana.

Methods S1 Chemical methods.

Table S1 List of primers used for the relative DNA copy number quantitation via qRT‐PCR.

Table S2 List of primers used for the relative RNA expression qRT‐PCR.

Please note: Wiley is not responsible for the content or functionality of any Supporting Information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

NPH-247-2601-s001.pdf (4.7MB, pdf)

Acknowledgements

We would like to thank Prof. E. Shani for providing Arabidopsis thaliana (Columbia ecotype) cell culture and Arabidopsis thaliana p35S::H2B‐RFP 2 transgenic line; Prof. A. Avni for providing Nicotiana benthamiana plants and maintaining the Arabidopsis thaliana cell culture; Prof. N. Ohad for providing Lamium amplexicaule seeds and Prof. N. Sade for providing Aegilops longissimi and Setaria viridis seeds. We thank Prof. L. Pizarro for her confocal training. We also want to thank Dr. A. Kessel for his help preparing Staphylococcus aureus (2XCS) and A. thaliana GyrA and GyrB DNA sequences for computational modelling and H. Failayev for her help in optimizing the models' visuals. We thank Dr. V. Holdengreber at the Rosalie and Harold Rae Brown Cancer Research Core Facility for the TEM microscopy operation and sample preparation; Dr. G. Meshulam for her PlantScreen™ Phenotyping System training and assistance and graduate student S. Levy for assistance in general lab work. This work was supported by the Israel Science Foundation (grant no. 1057/21 to RW). SL thanks the ADAMA Center for Novel Delivery Systems in Crop Protection, Tel Aviv University, for the financial support.

Data availability

All data supporting the findings of this study are available within the paper and its Supplementary Information. The Supporting Information file includes supporting figures and tables, general chemical methods, synthetic methods and chemical characterization methods and data.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Fig. S1 Nontargeted fluorophores do not specifically accumulate in mitochondria.

Fig. S2 FCCP disrupts mitochondrial accumulation of 3.

Fig. S3 Fluorescently labelled TPP localizes to mitochondria in various plant species.

Fig. S4 Fluorescently labelled TPP localizes to mitochondria in multiple organs of Aegilops longissimi.

Fig. S5 TPP moiety does not inhibit bacterial DNA gyrase supercoiling activity in vitro.

Fig. S6 CFX–TPP conjugate inhibits Arabidopsis mitochondrial DNA gyrase in planta.

Fig. S7 TPP moiety does not show phenotypic effects in Arabidopsis thaliana.

Methods S1 Chemical methods.

Table S1 List of primers used for the relative DNA copy number quantitation via qRT‐PCR.

Table S2 List of primers used for the relative RNA expression qRT‐PCR.

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Data Availability Statement

All data supporting the findings of this study are available within the paper and its Supplementary Information. The Supporting Information file includes supporting figures and tables, general chemical methods, synthetic methods and chemical characterization methods and data.

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