Plant Physiology and Biochemistry 103 (2016) 84e95
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Research article
Toxicity of canavanine in tomato (Solanum lycopersicum L.) roots is due to alterations in RNS, ROS and auxin levels Urszula Krasuska a, Olga Andrzejczak a, Paweł Staszek a, Wojciech Borucki b, Agnieszka Gniazdowska a, * a b
Department of Plant Physiology, Poland Department of Botany, Warsaw University of Life ScienceseSGGW, Nowoursynowska Str. 159, 02-776 Warsaw, Poland
a r t i c l e i n f o
a b s t r a c t
Article history: Received 20 January 2016 Received in revised form 3 March 2016 Accepted 3 March 2016 Available online 5 March 2016
Canavanine (CAN) is non-proteinogenic aminoacid and a structural analog of arginine (Arg). Naturally, CAN occurs in legumes e.g. jack bean and is considered as a strong allelochemical. As a selective inhibitor of inducible nitric oxide synthase in mammalians, it could act as a modifier of nitric oxide (NO) concentration in plants. Modifications in the content of endogenous reactive nitrogen species (RNS) and reactive oxygen species (ROS) influence root structure and architecture, being also under hormonal control. The aim of the work was to investigate regulation of root growth in tomato (Solanum lyco_ persicum L. cv. Malinowy Ozarowski) seedling by application of CAN at concentration (10 and 50 mM) leading to 50% or 100% restriction of root elongation. CAN at higher concentration led to slight DNA fragmentation, increased total RNA and protein level. Decline in total respiration rate after CAN supplementation was not associated with enhanced membrane permeability. Malformations in root morphology (shorter and thicker roots, limited number of lateral roots) were accompanied by modification in NO and ONOO localization; determined mainly in peridermal cells and some border cells. Although, CAN resulted in low RNS production, addition of exogenous NO by usage of NO donors did not reverse its negative effect, nor recovery effect was detected after roots imbibition in Arg. To build up a comprehensive view on mode of action of CAN as root growth inhibitor, it was shown an elevated level of auxin. To summarize, we demonstrated several secondary mode of action of CAN, indicating its toxicity in plants linked to restriction in RNS formation accompanied by simultaneous overaccumulation of ROS. © 2016 Elsevier Masson SAS. All rights reserved.
Keywords: Canavanine Cell viability DNA fragmentation Non-protein amino acid Phytotoxicity ROS RNS
1. Introduction Secondary metabolites play important role in plants as a
Abbreviations: ABA, abscisic acid; Arg, arginine; APF, 30 -(p-aminophenyl) fluorescein; BHT, butylated hydroxytoluene; CAN, canavanine; CTAB, cetyltrimethylammonium bromide; cPTIO, 2-(4-Carboxyphenyl)-4,4,5,5tetramethylimidazoline-1-oxyl-3-oxide; DAB, 3,3'-diaminobenzidine; DAF-FM DA, 0 0 4-amino-5-methylamino-2 ,7 -difluorofluorescein diacetate; GA, gibberellic acid; IAA, idolile-3-acetic acid; NBT, nitroblue tetrazolium; NO, nitric oxide; NOS, nitric oxide synthase; NPAA, non-proteinogenic amino acid; O 2 , superoxide anion; ONOO, peroxynitrite; PBS, phosphate buffered saline; ROS, reactive oxygen species; RNS, reactive nitrogen species; SNAP, S-nitroso-N-acetylpenicylamine; SNP, sodium nitroprusside. * Corresponding author. Warsaw University of Life Sciences-SGGW, Nowousynowska 159, 02-776 Warsaw, Poland. E-mail addresses:
[email protected] (U. Krasuska), andrzejczakolga5@ gmail.com (O. Andrzejczak),
[email protected] (P. Staszek), wojciech_
[email protected] (W. Borucki),
[email protected] (A. Gniazdowska). http://dx.doi.org/10.1016/j.plaphy.2016.03.005 0981-9428/© 2016 Elsevier Masson SAS. All rights reserved.
weapon against herbivores, microbes and/or competing plants. They also function as signaling compounds to attract animals (Weston and Duke, 2003). In spite of the name, chemicals belonging to the group of secondary metabolites are critical for plant survival and reproductive fitness and have been subjected to natural selection during evolution, resulting in their wide distribution among plant families (Wink, 2003). Non-proteinogenic amino acids (NPAAs) are examples of bioactive secondary metabolites that can be considered as useful taxonomic markers e.g. in Fabacea. NPAAs are defined as simple amino acids not included in protein structure. In general, they serve two important roles in higher plants, as key defense compounds and as mobile nitrogen storage reservoirs for tissues requiring nitrogen sources e.g. seeds (Wink, 2003). There are many endogenous NPAAs: L-canavanine (CAN), mimosine, albizziine, lathyrine, meta-tyrosine, 5hydroxytryptophan that are suspected to act as insect antifeedants, anti-microbials, or toxins responsible for plants' high
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allelopathic potential (Wink, 2003). CAN (2-amino-4guanidinoxybutanoic acid) is found in legumes, such as jack bean (Canavalia ensiformis (L.) DC.) (reviewed by Vranova et al., 2011). Jack bean has been shown to inhibit growth of many other neighboring plants. Synthesis of CAN is connected with light, photosynthetically active tissues are its production site, as was reported for green callus of Canavalia spp. (Hwang et al., 1996). CAN is structurally similar to arginine (Arg), and is considered as the guanidinooxy structural analog of Arg. Therefore, CAN is likely to exert some effects on Arg metabolism and other related amino acids. Rosenthal (1990) reported that in insects CAN was incorporated into proteins instead of Arg, as this amino acid is a substrate for arginyl-tRNA synthetase. Dysfunction of proteins with CAN incorporated instead of Arg is connected with decrease of basicity (Rosenthal and Harper, 1996). In insects, production of proteins containing CAN affected developmental processes and contributed significantly to expression of CAN's potent antimetabolic properties. These properties were also indicated in viruses, bacteria and fungi, and corresponded to disruption in DNA and RNA synthesis (Ekanayake et al., 2007). Growth of roots and its architecture is under control of phytohormones, mainly auxins. Indole-3-acetic acid (IAA) distribution in plants is connected with its polar (cell-to-cell) transport. Root architecture and IAA polar transport are also governed by reactive oxygen (ROS) and reactive nitrogen species (RNS) (Yu et al., 2014). Cross-talk of RNS/ROS and auxins, occurs in an model functional root system and is involved in regulation of development of adventitious roots, formation of lateral roots, growth of root hairs, and gravitropic response (review by Corpas and Barroso, 2015; Krasuska and Gniazdowska, 2015; Yu et al., 2014). It was observed for certain plants that phytotoxins inhibiting root growth may also induce oxidative stress manifested as overproduction of ROS and alterations in antioxidant enzymatic and non-enzymatic cellular system (for review see Gniazdowska et al., 2015). RNS include nitric oxide (NO) and peroxynitrite (ONOO) - the product of reaction of NO and superoxide anion (O-2). NO biosynthesis in plant cells occurs via several pathways which are classified into the oxidative and reductive ones. Reductive pathways depend on availability of nitrite (NO 2 ), while oxidative pathways require Arg as a substrate (for review see Gupta and Igamberdiev, 2015). It is also considered the existence of nitric oxide synthase like (NOSlike) enzyme, activity of which was detected mainly in peroxisomes, but also mitochondria and chloroplasts (Corpas et al., 2009). NOS activity depends on Arg and, as reported for mammalian, inducible isoforms of NOS (iNOS) is selectively inhibited by CAN (Abd El-Gawad and Khalifa, 2001). Thus, we suspect a close connection between CAN toxicity on root growth of tomato plants and modifications in NO tissue level. In the present study, the involvement of RNS and ROS was evaluated during CAN-induced restriction in growth of tomato roots. To investigate mode of action of CAN in plants we measured RNS (NO and ONOO) localization in roots, mainly in root tips. We have also determined concentration of NO 2 to find correlation of CAN treatment to reductive pathway of NO biosynthesis. Moreover, we focused on superoxide anion (O2 ) and hydrogen peroxide (H2O2) localization in roots of plants subjected to CAN at concentration leading to 50% inhibition of elongation growth or at concentration restricting root growth in 100%, as it is known that both ROS compounds influence root architecture in a different manner (Tsukagoshi et al., 2010). This RNS/ROS view was enriched by measurement of IAA content in roots. Moreover, to characterize harmful action of CAN during restriction of growth of tomato roots we investigated impact of tested NPAA on: total root respiration, membrane permeability, total protein and RNA level and DNA fragmentation. Data presented in the work broaden our knowledge on mode of action of potential
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allelopathic compounds; as so far, although ROS action in phytotoxicity of many allelochemicals is obvious (review by Gniazdowska et al., 2015), information on RNS involvement in their phytotoxicity are rare and very limited. 2. Material and methods 2.1. Plant material _ Tomato (Solanum lycopersicum L. cv. Malinowy Ozarowski) seeds were germinated in water in darkness at 20 C for 3 days. After radicle protrusion (day 0) seedlings of equal roots length (5 mm) were transferred to Petri dishes (Ø 15 cm) containing filter paper moistened with distilled water (control) or CAN (SigmaeAldrich) aqueous solutions (10, 50, 250 mM) (Fig. 1A). Dishes were carried in a growth chamber with 23/20 C, 12/12 h day/night regime for 3 days. Length of roots of seedlings was determined after 24 and 72 h of culture. The concentrations of CAN required for reduction of root length to 50% of the control, was accepted as IC50. The concentration, which totally inhibited elongation growth of roots, was accepted as IC100. 2.2. Germination test Tomato seeds were placed (25 per dish) on Petri dishes (Ø 9 cm), filled with filter paper moistened with distilled water (control) or CAN aqueous solutions (50, 250, 500 mM). Culture was carried for 4 days in darkness in 20 C. Seeds were considered germinated if the radicle had emerged through the seed coat. Experiments were repeated three times. 2.3. NO and cPTIO treatment of seedlings exposed to CAN To investigate the consequence of exogenous NO application on physiological effect observed after CAN treatment two different experiments were performed (Fig. 1). Single NO fumigation at the time point “0”, before exposition to CAN: tomato seedlings (developed from seeds imbibed for 3 days in water), at the time point “0” were shortly (3 h) treated with NO donors: sodium nitroprusside (SNP, 0.25 and 0.50 mM) and Snitroso-N-acetylpenicylamine (SNAP, 0.25 and 0.50 mM) at light. Then seedlings were washed in distilled water and transferred to Petri dishes and cultured in distilled water or CAN (10, 50 mM) solutions for 72 h (Fig. 1C). Root length of seedlings was measured after 72 h of culture. In the other test (repetitive, double fumigation with SNP at the time of seedlings treatment with CAN), seedlings treated with CAN (10, 50 mM) or seedling placed in distilled water for 24 h were shortly (3 h) treated with SNP (0.25 or 0.50 mM) at light, and placed again in CAN solutions or water. SNP treatment was repeated after 24 h, and seedlings were placed again in CAN solutions or water (Fig. 1E). The length of roots of seedlings was measured twice, 24 h after first SNP application, and then 24 h after second SNP application, which corresponds to 48 h and 72 h of the trial, respectively. Additionally, to check the influence of NO scavenging on the effect of CAN on growth of tomato roots 2-(4-Carboxyphenyl)4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO, 800 mM) was used. Seedlings were treated simultaneously with CAN (10, 50 mM) and cPTIO for 72 h (Fig. 1D). Length of the roots of tomato plants was determined after 72 h of culture period. Experiments were repeated three times. 2.4. Test of recovery effect after CAN treatment After 24 h of CAN (10, 50 mM) treatment seedlings were
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Fig. 1. The scheme of experiments. Treatment of tomato seedlings with CAN (A). Recovery experiment (B). Single short term application of NO donors at the time point “0” (C). Simultaneous treatment of tomato seedlings with CAN and cPTIO (D) Repetitive treatment with SNP (E). Details of treatment as described in Material and Methods.
transferred to Petri dishes containing filter paper moistened with distilled water or Arg (10, 50 mM) water solution (Fig. 1B). Control seedlings were treated with CAN (10, 50 mM) throughout the experiment. Length of roots of seedling was measured before recovery experiment (after 24 h CAN treatment) and after 24 h and 48 h of recovery, corresponding to 48 and 72 h of culture period, respectively. Experiments were repeated three times.
2.6. Electrolyte leakage determination Electrolyte leakage from roots of tomato seedling treated with CAN (10, 50 mM) for 24 h and 72 h was measured with a conductivity meter Elmetron CPC-505 (Poland). Roots were isolated from seedling, shortly washed in distilled water, and placed in 10 ml of distilled water (miliQ) at room temperature in darkness. Conductivity in the medium was measured after 1 h incubation. Results are expressed as % of total leakage observed after boiling roots for 10 min. Measurements were done in three independent experiments with threeefour repetition in each.
2.5. Test of cell viability Viability of tomato root cells was determined using Evans blue staining. Whole control seedlings or seedlings treated with CAN (10, 50 mM) for 24 h and 72 h were incubated in 0.25% solution of Evans blue for 30 min at room temperature. Then seedlings were washed twice in distilled water and roots were isolated, weighed and homogenized in 1 ml of 1% solution of sodium dodecylsulphate (SDS). After 5 min centrifugation at 12 000 g at 4 C, supernatant was collected and absorbance was measured at 600 nm (Hitachi U2900 spectrophotometer). Three roots were used for one repetition. Data are means of three measurements from each three to four sets of experiments.
2.7. Measurement of oxygen consumption Oxygen consumption by roots of tomato seedlings was measured using Clark type oxygen electrode (Oxygraph 23107, Hansatech). After isolation, roots of tomato seedling were placed in distilled water in darkness for 10 min. Then, roots (of approximately 15e20 seedlings) were transferred to measurement chamber. Assays were performed for 5e10 min at temperature 25 C, in darkness, and at atmospheric CO2 concentration, in three-four repetition. Oxygen consumption by roots was expressed as nmol O2 min1g1FW.
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2.8. Imaging of endogenous NO and ONOO- in root tissue
2.11. In situ H2O2 localization
NO and ONOO detection were done by confocal laser scanning microscopy (CLMS) Leica TCS SP5II (Leica Microsystems CMS, Wetzlar, Germany) equipped with an acousto-optical beam splitter (AOBS) and an upright microscope stand (DMI 6000), using 4amino-5-methylamino-20 ,70 -difluorofluorescein diacetate (DAFFM DA, Invitrogen) and 30 -(p-aminophenyl) fluorescein (APF, Invitrogen) markers (respectively). The confocal pinhole diameter was automatically set to so called ‘1 Airy’ unit to reduce the effect of light diffraction on image formation. As a result, a good compromise between signal-to-noise ratio and resolution were achieved. Root autofluorescence had been reduced by an appropriate technical mean (gain adjustment) before the fluorescence dye was administrated. The resulted autofluorescence level was negligible or not detectable. Roots of tomato seedlings grown in water (at time point 0, 24 h and 72 h) or in CAN (10, 50 mM) (at time point 24 h and 72 h) were isolated and shortly washed in distilled water. Roots were incubated in the presence of DAF-FM DA (10 mM) for NO detection or APF (20 mM) for ONOO- detection in darkness at room temperature. Observations were done using excitation/emission range of 496 nm/510e530 nm (for DAF-FM DA) and 488 nm/505e525 nm (for APF). Parameters for fluorescence microscopy were identical for all experiments. All images were processed in the same manner using the ImageJ program. At least ten roots were tested under each experimental condition and five independent repeats were analyzed. The representative images for control and Can treated plants at appropriate time points of experiments are shown.
Localization of H2O2 was done with 3,3'-diaminobenzidine (DAB) staining. Roots of control seedlings at the beginning of the experiment (0 h), after 24 and 72 h of culture or treated for 24 and 72 h with CAN (10, 50 mM) were washed twice in distilled water and incubated with DAB solution (1 mg ml1) with 0.05% Tween-20 and 2 mM DMSO. Staining was done for 4 h at room temperature, in darkness. Experiments were done in five independent repetitions.
2.9. Measurement of NO 2 concentration Total nitrite concentration in roots of tomato seedlings was determined spectrophotometrically with modified Griess reagent (Sigma G4410). Roots (30 mg of fresh weight) isolated from control seedlings at the beginning of treatment (0 h), after 24 h 72 h of culture or treated for 24 and 72 h with CAN (10, 50 mM) were washed twice in distilled water and homogenized in 0.5 ml of 0.1 M Tris-HCl (pH 7.0) buffer with 2% polyvinylpolypyrrolidone (PVPP). Homogenates were centrifuged at 12 000 g for 10 min at 4 C. Supernatants (100 ml) were transferred to appropriate wells of 96well microplate filled with 100 ml modified Griess reagent. After 15 min incubation in darkness at room temperature NO 2 concentration was measured at 540 nm using microplate absorbance reader (Sunrise, Tecan, Switzerland) and compared to standard curve with sodium nitrite, used as a standard. Concentration of nitrite was expressed as pmol mg1 FW. The data are means of three measurements from each of three sets of experiments. 2.10. In situ O2 localization Localization of O2 was performed using method of nitroblue tetrazolium (NBT) staining. Roots isolated from control seedlings at the beginning of the experiment (0 h), after 24 h and 72 h of culture or treated for 24 h and 72 h with CAN (10, 50 mM) were washed twice in distilled water and incubated for 20 min in darkness at room temperature in 2 mM NBT (SigmaeAldrich), freshly prepared in 10 mM Tris-HCl buffer pH 7.4 with addition of 2 mM DMSO. O2 . was visualized as a deposit of dark blue insoluble formazan compound. Additionally roots of seedlings treated with SNP and cPTIO (as described above) were incubated with 2 mM NBT in 10 mM TrisHCl buffer pH 7.4 with addition of 2 mM DMSO. After 20 min of incubation in darkness O-.2 visualization was done. Experiments were done in five independent repetitions.
2.12. RNA isolation Total RNA was isolated according to Chomczynski and Sacchi (1987). Tomato seedlings of control and treated plants were briefly washed in distilled water, then dried on filter paper, and roots after isolation were weighted (to obtain samples of 150 mg) and frozen in liquid nitrogen. Homogenized tissues were mixed with the solution of 4 M guanidine thiocyanate, 0.025 M sodium citrate, 0.5% (w/v) N-laurosylsarcosine (Sarkosyl), 2 M sodium acetate pH 4.0 and water-saturated phenol at 1:1 (v/v). Then RNA was precipitated by addition of the salt-mix (0.8 M sodium acetate and 1 M sodium chloride), and isopropanol at 1:1 ratio, and extracted using chloroform and isoamyl alcohol (49:1). Concentration of total RNA, after ethanol-wash step and dilution in water was determined using NanoDrop 2000 (Thermo Scientific) spectrophotometer at 260 nm. The obtained RNA quality was calculated after absorbance measurement at 260/230 nm and 260/280 nm. RNA electrophoresis (1 ml of total RNA) was done using 1.5% agarose gel with addition of ethidium bromide (1 mg ml1), and typical electrophoretic pattern was shown. The results were also calculated and presented as mg RNA mg1 FW. Three - four independent repetitions were done. 2.13. DNA isolation and analyses of DNA fragmentation DNA was isolated according to Murray and Thompson (1980). Isolated roots of control and CAN treated tomato seedlings were weighted (to obtain samples of 150 mg), then grounded in liquid nitrogen to a fine powder. Samples were incubated in extraction buffer: 0.1 M Tris-HCl pH 8.0, 10 mM EDTA, 0.7 M NaCl, 0.2% 2mercaptomethanol, and 2% cetyltrimethylammonium bromide (CTAB) for 30 min at 60 C, slightly shaking. After centrifugation (12 000 g, 5 min, 4 C), DNA was extracted with chloroform-isoamyl alcohol. Addition of RNase A enable to remove RNAs from samples. DNA was precipitated with isopropanol. Isolated DNA was resuspended in Tris-actetate EDTA buffer. Concentration of DNA was measured using NanoDrop 2000 (Thermo Scientific) spectrophotometer. DNA samples which measured absorbance (at 260/280 nm and 260/230 nm) values greater than 1.8 were used for further analysis. One ml of isolated DNA was electrophoretically separated on 1% agarose gel with ethidium bromide. Three independent repetitions were done. 2.14. Total soluble protein extraction Protein concentration measurement was performed using Bradford reagent. Tomato seedlings roots were separated and rinsed in distilled water. After fresh weight determination, roots were homogenized in 50 mM Tris-HCl pH 7.0, 2% (w/v) polyvinylpolypyrrolidone (PVPP), 10% (v/v) glycerol, 5 mM dithiothreitol (DTT), 1% (v/v) cocktail of protease inhibitors (Sigma) in ice bath. After centrifugation 12,000 g for 10 min at 4 C, supernatant was collected, and protein content was measured at 595 nm using microplate reader (Sunrise, Tecan). The results were presented as mg mg1FW. The experiment was done in three independent
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repetitions. 2.15. Indole-3-acetic acid (IAA) determination IAA determination was performed by ELISA method according to Marcussen et al. (1989) and Elsorra et al. (2004) with some modifications. 2.15.1. Sample preparation Samples (0.2 g of roots isolated from control seedlings after 24 and 72 h of culture or seedlings treated with CAN (10, 50 mM) after 24 and 72 h) were homogenized in 80% methanol with butylated hydroxytoluene (BHT, 20 mg L1) and incubated for 24 h at 70 C. After incubation samples were centrifuged at 10 000 g for 15 min, and supernatants were collected. The remain pellets were washed with 80% methanol with BHT, and after centrifugation supernatants were added to the first lots. At the same time standard solution of IAA in 80% methanol with BHT was prepared. Samples and IAA standards were methylated: 2.0 M (trimethylsilyl)diazomethane in hexanes (Sigma 362832) for 30 min at room temperature. After methylation step, the reaction was stopped by adding 5 ml of 0.05 M acetic acid. Samples were left to evaporate. Afterwards pellets were resuspended in phosphate buffered saline (PBS). Methylated IAA (Sigma I9770) was used as a positive control. 2.15.2. Microplates preparation Microplates were covered with antibodies: Rabbit Anti-Mouse IgG (RAMIG Sigma) (25 mg ml1 in 0.05 M NaHCO3 buffer, pH 9.6) overnight at 4 C. Next day, antibodies were removed, and plates were covered with monoclonal Anti-Auxin (Sigma, A0855) (1 mg ml1 in 50 mM TBS buffer pH 7.8). Incubation was performed overnight at 4 C. After removal of antibodies plates were washed twice with deionized water, and then samples, IAA standards, and positive control were added (50 ml per well) and incubated in TBS buffer at 4 C for 1 h. Next, IAA tracer (Agrisera AS09517) (3 ml per 5 ml in TBS buffer with 0.1% gelatin) was added to each well and incubation was performed for 1 h at 25 C. Plates were washed twice with deionized water and incubated at 37 C with p-nitrophenylphosphate disodium (pNPP) at concentration 1 mg ml1 in 1 M diethanolamine (DEA) solution with 0.5 mM MgCl2, as long as color was observed. The reaction was stopped with 5 ml 5 M KOH, and absorbance was read at 405 nm with referential wave 605 nm in microplate reader (Sunrise, Tecan). Concentration of IAA was expressed as pmol mg1 FW. Experiment was done in 4e5 independent repetitions. 2.16. Statistics Data were analyzed using the Statistica Software. For experiments, distribution of the data was checked using ShapiroeWilk test. Mean differences were calculated using Student's t-test and homogenous groups were evaluated using Tukey's HSD post-hoc test. Standard errors (SE) were also provided to indicate the variations associated with the particular mean values. 3. Results
after 3 days of culture (Table 1), but the effect was not stable, as after 4 days seeds imbibed in 500 mM CAN germinated in 90%, at the rate similar to the control.
3.2. Elongation growth of tomato roots is inhibited by CAN Roots of control tomato plants grow well in water (Fig. 2, Table 2). After 24 h their length was about 40 mm, the prolonged culture led to elongation of the roots to about 90 mm. CAN visibly repressed root growth. At lowest concentration (10 mM) CAN inhibited root growth in 50%, while at concentration of 50 or 250 mM growth of tomato roots was totally restricted (Fig. 2), so significant differences in root length were noticed in plants exposed to 50 or 250 mM CAN for 24 or 72 h (Fig. 2). Thus, we have selected 50 mM concentration of CAN as IC100 and 10 mM as IC 50. CAN in these two chosen concentrations was used in further experiments.
3.3. Toxic effect of CAN on roots of tomato seedlings is not reversible Recovery effect of CAN was investigated using Arg, as CAN is considered as antimetabolite of Arg. Arg did not affected growth of tomato seedlings (Table 2). Length of roots of seedlings imbibed in Arg (10, 50 mM) was the same as length of roots of plants cultured in water. Transfer of seedlings from CAN solution into Arg of appropriate concentration or water did not removed inhibitory effect of CAN on tomato root growth (Table 2).
3.4. CAN does not lower viability of tomato roots Viability of roots of plants imbibed in CAN solution was determined by Evans blue staining (Fig. 3). Only slight, statistically insignificant decline (of 7e8%) of viability was detected in tomato roots treated with both 10 and 50 mM CAN after 72 h. Short term treatment of plants with CAN did not alter cell viability, as compared to control.
3.5. CAN only at high concentration leads to slight DNA fragmentation in cells of tomato roots The integrity of DNA in cells of tomato roots was monitored after 24 and 72 h of CAN exposure. DNA fragmentation from control tomato roots, showed 1 band with a very little smear only after 72 h of culture. Fragmentation of DNA from tomato roots treated with CAN was not observable after 24 h, slightly more visible smear was noticed after 72 h of the culture, with no difference depending on NPAA concentration (Fig. 4).
Table 1 Germination rate of tomato seeds imbibed in water (control) and in water solution of CAN at concentration 50 mM, 250 mM and 500 mM after 3 or 4 days. Plant treatment
Germination (%)
Control (water) CAN (50 mM) CAN (250 mM) CAN (500 mM)
58 52 56 38
3.1. Germination of tomato seeds is not altered by CAN treatment Tomato seeds germinated well after imbibition in water. Three days long imbibition resulted in 58% germination, while after next 24 h more than 85% of seeds were germinated (Table 1). CAN at lower concentration (50 and 250 mM) did not alter tomato seed germination, both after 3 and 4 days of imbibiton. A slight inhibition of tomato seeds germination was observed only in CAN 500 mM
3 days ± ± ± ±
3 8 8 8*
72 h 85 76 83 88
± ± ± ±
6 4 9 6
Values are average ± SE of 3 replicated experiments. Asterisk (*) indicate significance from control at the same time of culture period at P 0.05 based on Student's tests.
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Fig. 2. Tomato seedlings grown for 24 or 72 h in water (control) or CAN aqueous solutions (10, 50, 250 mM). Images of representative seedlings are presented.
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twice higher than length of roots of plants treated with 10 mM CAN alone. Stimulatory effect of cPTIO was much less pronounced in roots of seedlings exposed to 50 mM CAN (Fig. 5). In contrast, a single (one-time) short term pre-treatment of seedlings with NO donors had no positive consequence on elongation of the roots. Both SNP or SNAP short term application at the beginning of the experiment led to slight inhibition in growth of root of plants cultured in water and 10 mM CAN. No effect of short term fumigation of seedlings with NO donors at time point “0” was detected in roots of seedlings imbibed in 50 mM CAN (Fig. 5). Application of SNP (at higher concentration - 500 mM) in two intervals (after 24 h and 48 h after starting the experiment) resulted in 32% inhibition of root growth of control seedlings. SNP at lower concentration (250 mM) had less effect on growth of roots of control plants, it was inhibited only in about 15% (Fig. 6). Double application of SNP (independently of SNP concentration) did not alter growth of roots of seedlings imbibed in 50 mM CAN (Fig. 6). Similarly, no influence of repetitive SNP fumigation was
Table 2 Length (mm) of roots of tomato seedlings cultured in water (control) or treated with CAN (10 or 50 mM) or after recovery test consisting of: removal from CAN solution and imbibition in Arg (10 or 50 mM) or imbibition in water. Length of roots was determined at the beginning of recovery experiment (Fig. 1B) and after 24 and 72 h of culture. Treatment
Time of experiment (h)
CAN 10 mM
CAN 50 mM
Arg 10 mM
Arg 50 mM
H2O
0
24
e þ
e
e
e
þ
18 ± 2.4c
43 ± 4.2b
e e
13.5 ± 2.1 7.1 ± 0.6e
d
e
e þ
e e
e e
e
e
þ
e
e
20.2 ± 1.7
e þ þ
e
e
þ
e
18.3 ± 2.4c d
c
72 87.9 ± 11.6a bc
25.6 ± 6.5 7.1 ± 0.5 e
46.9 ± 12.4b 7.5 ± 0.6e
42.8 ± 3.6b
88.0 ± 10.9a
39.2 ± 10.8b
74.4 ± 13.1a
c
e
e e þ
þ e þ
e e e
e þ e
13.1 ± 1.4 10.1 ± 1.9d 7.2 ± 0.8e
23.4 ± 5.8 15.6 ± 1.8c 7.2 ± 0.7e
43.3 ± 12.5b 36.6 ± 10.3b 7.7 ± 0.7e
e
þ
e
e
þ
7.8 ± 0.8e
8.2 ± 1.2
8.8 ± 1.3de
Values are average ± SE of 3 replicated experiments; a,b, … - homogenous groups.
3.6. CAN does not increase membrane permeability Electrolyte leakage from cells of roots of control seedlings did not alter as the culture period was prolonged, and was lower than 20% (Table 3). CAN did not influence membrane permeability. Its value in treated seedlings was stable during the experiments and comparable to control, did not exceed 21%.
noticed in plants treated with 10 mM CAN. Stimulation of elongation growth of roots by repetitive SNP application was observed only after higher concentration of SNP (500 mM) in seedlings growing in 10 mM CAN. Roots of these plants were 35% longer as compared to roots of plants exposed to 10 mM CAN only. 3.9. CAN inhibits NO emission and transiently enhances ONOO production in tomato roots
3.7. CAN lowers oxygen consumption by tomato roots Total respiration rate by roots of control tomato seedlings did not differ significantly during culture period and varied from 7.3 to 6.5 nmol O2 min1g1FW after 24 or 72 h respectively (Table 3). CAN decreased total root respiration rate of tomato seedlings. Oxygen consumption by roots of seedlings treated with NPAA at higher (50 mM) concentration decreased two fold to around 2.7 nmol O2 min1g1FW, and remained at the same level till the end of experiment. CAN at lower (10 mM) concentration inhibited respiration rate of tomato roots in 30% or 46% after 24 or 72 h, respectively (Table 3). 3.8. NO donors and scavenger only slightly modify growth of roots of CAN treated plants Application of 800 mM cPTIO slightly inhibited growth of tomato roots in water (Fig. 5). Its addition to the medium used for culture of CAN treated plants resulted in beneficial effect on root elongation. Length of roots of seedlings treated with 10 mM CAN and cPTIO was
CAN treatment, especially in the concentration 50 mM, resulted in a substantial increase in root thickness. The production of the border cells by the root cups was also enhanced comparing to the control (Fig. 7a,b,c,d). The CLSM in vivo detection of NO and ONOO in roots of control and CAN-treated seedlings is presented at Fig. 7a,b,c,d. Generally, NO fluorescence was localized mainly in epidermis and border cells in control and CAN treated plants (Fig. 7a,b). A similar image was obtained for ONOO fluorescence (Fig. 7c,d). NO signal in CAN treated roots was generally reduced as compared to control, but was visibly brighter in border cells especially after 72 h. In case of control roots NO was localized mainly in the epidermis of the elongation zone. After CAN application an increase in NO related fluorescence was detected in the zone of cell division as well as root cups and border cells. ONOO related fluorescence was uniformly distributed along the root surface of both control and CAN-treated plants, independently of the CAN concentration. APF fluorescence in root apex was declining slightly as culture period was prolonged. In root tips of plants exposed to 50 mM CAN for 72 h a bright signal was noticed
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both in control and CAN treated roots. No CAN dependent modifications in NO 2 concentration in tomato roots were detected (Table 3). 3.11. CAN strengthened visualisation of O2 and H2O2 in roots of tomato plants
Fig. 3. Cell viability test by Evans blue staining of roots of tomato seedlings cultured in water or treated with CAN (10 or 50 mM) after 24 or 72 h of culture. Images of representative test are presented. Inserted Table includes average values ± SE of 3 replicated experiments. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
Superoxide radical (O2 ) detection was performed using NBT staining method (Fig. 8A). The levels of O2 staining were quite similar in roots of control plants and plants treated with 10 mM CAN after 24 h. However, the time-course revealed decline coloration in control roots and roots of plants treated with 10 mM CAN. In opposite, a strong blue staining appeared in roots treated with 50 mM CAN. This increase was more visible after 24 h than 72 h and characteristic for root apex. H2O2 detection was performed using DAB staining method, observed as brown precipitation of polymerized DAB (Fig. 8B). After 24 h of experiment initiation, control and CAN treated plantlets exhibited a much stronger level of staining than plantlets cultured for additional 48 h (Fig. 8B). Dark brown coloration was detected mainly in roots of CAN treated seedlings after 24 h of exposure to NPAA. No variation in H2O2 accumulation were observed in roots treated with 10 or 50 mM CAN. The most visible staining was detected in the root tips. Whereas, H2O2 content decreased visibly in control roots as culture period was prolonged, only light coloration was noticed in root tips (Fig. 8B). Additional NBT staining was done after plants simultaneous treatment with CAN and cPTIO or SNP. cPTIO led to decline coloration of the roots both of control and CAN treated seedlings. The combination of CAN and SNP resulted in only slight coloration of roots, but led to distinct visualization of formation of lateral roots, observed mainly after treatment with 10 mM CAN and in control plants. SNP declined blue coloration of roots of seedlings exposed to 50 mM CAN, and did not increase formation of lateral roots (Fig. 9). 3.12. CAN treatment increases total RNA concentration in tomato roots Content of total RNA in roots of tomato seedlings growing in water declined for 40% during 2 days of experiment (Fig. 10). CAN (50 mM) application resulted in elevated RNA concentration, observed mainly after short period (24 h). After prolonged (72 h) exposition to NPAA, higher than in control seedlings RNA content was observed only for 50 mM CAN (Fig. 10). 3.13. CAN slightly increases total proteins content in tomato roots
Fig. 4. Typical patterns of DNA fragmentation assay. DNA was isolated from roots of tomato seedlings cultured in water (control) or treated with CAN (10 or 50 mM) after 24 or 72 h of experiment, M- marker.
mainly in root caps (Fig. 7d). 3.10. CAN does not alter NO 2 concentration in roots of tomato seedlings Nitrite concentration declined as culture period was prolonged
Content of total proteins in roots of control tomato seedlings declined twice during experiment; it dropped from value of 4.45 mg mg1FW to 2.18 mg mg1FW (Fig. 10). CAN treatment led to increase of protein concentration as compared to control, observed mainly after short period of NPAA application. After prolonged exposition to CAN, higher than in control seedlings protein content was observed only for 50 mM CAN (Fig. 10). 3.14. Treatment with CAN enhances IAA concentration in roots of tomato seedlings IAA concentration in roots of control plants increased in 35% during two days of culture (Table 3). Treatment with CAN led to enhancement in IAA level immediately after CAN application and independently of CAN concentration. In roots of plants exposed to CAN IAA concentration was about 60e70% higher as compared to control plants (Table 3).
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Table 3 1 1 Electrolyte leakage (%), oxygen uptake (nmol min1mg1 FW), concentration of NO FW) in roots of control tomato seedlings (growing in 2 (nmol mg FW) and IAA (pmol mg water) and roots of tomato seedlings treated with CAN (10 and 50 mM) for 24 or 72 h. Plant treatment
Electrolyte leakage (%)
O2 uptake (nmol min1mg1 FW)
NO 2 concentration (pmol mg1FW)
IAA concentration (pmol mg1FW)
24 h
72 h
24 h
72 h
24 h
72 h
24 h
72 h
Control (water) CAN 10 mM CAN 50 mM
17 ± 3a 19 ± 3a 14 ± 2a
20 ± 4a 21 ± 4a 16 ± 3a
7.3 ± 1.3 5.1 ± 0.9 2.7 ± 0.4*
6.5 ± 1.4 3.5 ± 0.5* 2.8 ± 0.3*
70 ± 8a 65 ± 6a 62 ± 7a
30 ± 5b 32 ± 4b 31 ± 5b
28 ± 6 46 ± 10* 50 ± 10*
38 ± 7 60 ± 12* 58 ± 10*
Values are average ± SE of 3e4 independent experiments. Asterisk (*) indicate significance from control at the same time of culture period at P 0.05 based on Student's test; a,b-homogenous groups.
Fig. 5. Length of roots of tomato seedlings shortly (3 h) pre-treated with NO donors (SNP or SNAP) before culture in water or exposure to CAN (10, 50 mM) determined after 72 h of culture, and length of roots of seedlings treated with cPTIO (800 mM) or simultaneously with CAN (10, 50 mM) and cPTIO (800 mM). Values are presented as % of respective control: length of seedlings growing in water established as control to seedlings treated with SNP, SNAP or cPTIO; length of seedlings growing in 10/50 mM CAN established as control to seedlings treated simultaneously with 10 or 50 mM CAN and cPTIO, pre-treated with SNP or SNAP and then cultured in 10 or 50 mM CAN. Values are average ± SE of 3 replicated experiments. Asterisk (*) indicate significance from appropriate control at P 0.05.
4. Discussion CAN is a structural analogue of Arg, and its antagonist. The dmethylene group of Arg, in CAN is replaced by oxygen altering chemical and physical properties of the compound (Hwang et al., 1996). This NPAA is also known as an antimetabolite, toxic to herbivores e.g. tobacco hornworm (Manduca sexta L.) (Rosenthal, 1990). Toxicity of CAN is linked to its incorporation into protein structure, leading consequently to metabolism disruption (Rosenthal and Harper, 1996) and recognized as primary effect of its
Fig. 6. Length of roots of tomato seedlings after doubled fumigation with SNP (250, 500 mM) during treatment with CAN (10, 50 mM) determined after 48 and 72 h of treatment with NPAA. Values are presented as % of respective control: length of seedlings growing in water established as control to seedlings treated with 250 or 500 mM SNP; length of seedlings growing in 10/50 mM CAN established as control to seedlings treated with 10/50 mM CAN and 250 or 500 mM SNP, at time point 48 or 72 h respectively. Values are average ± SE of 3 replicated experiments. Asterisk (*) indicate significance from appropriate control at P 0.05.
action. CAN is metabolized to canaline and urea in reaction catalyzed by arginase (Jang et al., 2002). Canaline is toxic, and acts as inhibitor of activity of vitamin B6-containing enzymes. Moreover, just recently Kamo et al. (2015) had found that in plant tissues CAN may be converted to cyanamide - a strong phytotoxin and allelopathic compound (Soltys et al., 2011, 2012; 2014). This reaction is catalyzed by unidentified enzyme of high substrate specificity (Kamo et al., 2015). It is also known for mammalians, that characteristic properties of CAN make it a selective inhibitor of inducible nitric oxide synthase (iNOS) isoform. CAN declines also cellular nitrate level, as indicated in experimental endotoxemia (Abd ElGawad and Khalifa, 2001). CAN is stored in seeds of jack bean, thus is regarded as nitrogen source during seed germination (Rosenthal, 1990). In addition it is found in leaves of suderlandia (Sutherlandia frutescens (L.) R.Br.), the plant used often in alternative medicine, especially in south Africa (Colling et al., 2010).
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Fig. 7. NO-related DAF-FM fluorescence (a,b) and ONOO- related APF fluorescence (c,d) in root apex of tomato seedlings as affected by CAN (10 or 50 mM) treatment for 24 h (a,c) or 72 h (b,d). The capital letters A-D show the bright-field images corresponding to fluorescence images a-d. Bar ¼ 250 mm (a, b); bar ¼ 75 mm (c, d). Representative images are shown.
Natural occurrence of CAN in plant tissues points out on allelopathic properties of this NPAA. Although, our results indicate that CAN has no negative effect on germination of tomato seeds, probably due to its immobilization in seed coat. On the other hand, we have previously shown that imbibition of apple (Malus domestica Borkh.) embryos in CAN solution prevented germination. Apple embryos imbibed in CAN were deeply dormant likewise ones treated with abscisic acid (ABA, phytohormone responsible for maintenance of seed dormancy state) (Krasuska et al., 2014). CAN showed an inhibitory effect on elongation growth of rice (Oryza sativa L.) seedlings, similarly to ABA, both in the presence and absence of gibberelic acid (GA) (Nakajima et al., 2001), suggesting that the mode of action of CAN is not due to inhibition of GA synthesis. In our experiments, toxicity of CAN was observed during post-germination growth of tomato seedlings, when embryonic roots ruptured seeds coats. CAN inhibited tomato root growth in dose-dependent manner. This effect was not reversible after removal of CAN from growing medium, even if seedling were supplemented with Arg (no recovery effect). These findings are in agreement with inhibition of root growth of tomato seedlings after treatment with cyanamide (Soltys et al., 2012). The authors concluded that prolonged culture of tomato seedlings in cyanamide caused irreversible alterations in plant metabolism. Similar effect of cyanamide was observed in onion (Allium cepa L.) roots; only short term treatment with this allelochemical was fully reversed by transfer of bulbs into distilled water (Soltys et al., 2011). Therefore, taking together information on cyanamide phytotoxicity, and data suggesting direct link of CAN and cyanamide biosynthesis (Kamo et al., 2015) we could partially explain CAN negative impact on
living organisms, noticed even at low (mM) NPAA concentration. Although, CAN (50 mM) application for 72 h strongly and irreversibly inhibited growth of tomato roots, no lethal effect of this NPAA was observed. High cell viability, detected by Evans blue staining, was also confirmed by measurement of membrane permeability, values of which were comparable to the control. Such reaction to allelochamicals is uncommon, as most phytotoxins or their mixtures act rather as enhancers of membrane deteriorations by induction of oxidative stress (Gniazdowska et al., 2015). Knowing that CAN may influence nucleic acids synthesis (Ekanayake et al., 2007) and suspecting that DNA degradation could be linked to stress-induced root growth inhibition, we analyzed DNA fragmentation in root cells. Short term exposition (24 h) to CAN did not lead to DNA degradation in roots of tomato seedlings, but prolonged treatment with 50 mM NPAA resulted in slight DNA degradation, suggesting potentially harmful effect of CAN toward DNA. In human acute leukemia Jurkat T cells treatment with this NPAA led to apoptosis, accompanied by DNA fragmentation (Jang et al., 2002). ROS impact membrane permeability, but H2O2 participates also in stiffening process, leading to cross-linking of cell wall polymers (Müller et al., 2009), thus we are allowed to propose that CAN could influences cell wall thickness rather than membrane properties. Our data indicate modifications in H2O2 localization in roots of seedlings treated with CAN (particularly 50 mM). The most intensive and visible coloration pointing on H2O2 accumulation was detected mainly in the root tips. This is probably linked to structural malformations of root tips. High content of H2O2 is usually connected with disruption of root hair formation,
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Fig. 8. Visualization of superoxide radical (O2 ) detected by NBT (A) and hydrogen peroxide (H2O2) detected by DAB staining. Detections have been done on roots of control tomato plants, growing in water, 24 and 72 h after starting the experiment and on roots of treated seedlings 24 or 72 h after imbibition in CAN solution (10 or 50 mM). A representative results of 4e5 repetitions are shown.
decline of root growth and shortened root meristem (Tsukagoshi et al., 2010). Rapid increase in H2O2 level is typical reaction after
application of most allelochemicals; e.g. was noticed in Arabidopsis roots treated with p-hydroxybenzoic acid (Guan et al., 2014). Similarly, cyanamide treatment of onion or corn (Zea mays L.) roots caused over accumulation of H2O2 and was linked to disruption of root meristem (Soltys et al., 2011, 2014). It was proposed that cyanamide via disturbances in ROS level causes transfer
Fig. 9. Visualization of superoxide radical (O-.2 ) detected by NBT staining done on roots of control tomato plants 72 h after starting the experiment and on roots of CAN treated seedlings 72 h after imbibition in CAN solution (10 or 50 mM) and simultaneous application of cPTIO (250 mM) or SNP (500 mM). A representative results of 4e5 repetitions are shown.
Fig. 10. Total proteins and RNA content in roots of control tomato plants, growing in water, 24 and 72 h after starting the experiment and in roots of treated seedlings after 24 or 72 h imbibition in CAN solution (10 or 50 mM). Typical electrophoretic patterns of isolated RNA are shown. Values are average ± SE of 3 replicated experiments. Asterisk (*) indicate significance from appropriate control at P 0.05.
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of the transition zone from cellular proliferation to differentiation. An increase of O2 content in elongation and differentiation zones of roots of onion bulbs was observed at the beginning of cyanamide treatment, and after prolonged exposure to phytotoxin also in root tips (Soltys et al., 2011). Treatment of tomato seedling with CAN in addition to elevated H2O2 level led to over accumulation of O2 . As enhancement of production of both compounds (H2O2 and O2 ) was not lethal, and did not influence membrane integrity or DNA degradation we can assume that location of ROS burst is rather in apoplastic space, and could refer to cell wall structure modification e.g. lignification depending on ROS and RNS level. Thus, it is suggested that modifications of cell wall properties could be of significant, but secondary effect of CAN action, and need more interest in future research. Quite recently, it was also shown that NO participate in the lignin biosynthesis (Corti n et al., 2014). Into RNS family there are included NO (at Monzo different redox state), but also ONOO and S-nitrosothiols (SNOs). Accumulation of ONOO can be considered as an indicator of simultaneous NO and O2 overproduction because it is the product of NO and O2 reaction. Exposure of Arabidopsis seedlings to phydroxybenzoic acid induced NO emission, reversed by inhibitors of NOS and nitrate reductase activity (Guan et al., 2014). We demonstrated that CAN treatment of tomato seedlings modified NO and ONOO signals observed under CLSM. Moreover, treated roots were characterized by production and secretion of mucilage, acting probably as a response to stress. CAN stimulated also formation and release of border cells. Relationships between physiological quality of border cells and root growth are not very obvious and most likely depend on the examined phytotoxin (Soltys et al., 2014). After CAN treatment border cells were characterized by high fluorescence signal (detected as DAF-FM fluorescence) corresponding to enhancement in NO emission. It cannot be ruled out, that observed reaction is due to alteration in border cell viability, e.g. due to CAN incorporation into some proteins but it needs more detailed study. NO 2 acts as an important plant substrate for NO formation, particularly via nitrate reductase dependent pathway, therefore we have focused on its concentration in roots of plants treated with CAN. NO 2 concentration after CAN application was similar to the control. These findings let us to consider that elusive NOS-like activity and existence could be important for intact root growth. At this point of view, we may suggest that CAN (per se) probably do not influence reductive pathway of NO biosynthesis, but rather its oxidative one. All the more, in plants exposed to both CAN concentration, decrease of NO 2 was observed after 72 h of the culture. These results correspond to the ones described by Negi et al. (2010). In tomato mutants (shr), characterized by extremely short roots, application of NO scavengers or inhibitors of NOS-like activity restored root elongation growth. The authors indicated that in shr mutants nitrate reductase pathway of NO biosynthesis did not play an important role (Negi et al., 2010). Thus, on the basis of these data, we may suppose that CAN function as a potent stress factor lowering NO level below the concentration set in hormetic window or “nitrosative door”. NO treatment increases number of root hairs and promotes their elongation (Lombardo et al., 2006), as was proven by n et al., 2014). application of NO scavengers, e.g. cPTIO (Corti Monzo Addition of cPTIO during CAN treatment (at lower concentration) resulted in stimulation of elongation root growth, but decreased number of lateral root primordia. However, application of cPTIO did not reverse, nor strengthened inhibitory effect of CAN at higher concentration. On the other hand, short-term treatment of tomato seedlings with NO donors and further culture in medium containing CAN stimulated formation of lateral root primordia, observed particularly at lower concentration of this NPAA. Inhibitory effect of CAN at 10 mM concentration was partially reversed by repetitive
short-term (3 h) application of NO donor. These findings points at CAN as a potent modulator of RNS metabolism/biosynthesis, and confirm its role of strong inhibitor of root growth. Respiration rate reflects total metabolic activity of the tissue, thus characterize general physiological condition of the cells. Same allelochemicals are known to act as nonspecific inhibitors of mitochondrial respiration. Higher concentration of a-pinene decreased oxygen consumption in isolated mitochondria of maize coleoptiles and primary roots (Abrahim et al., 2003). The authors pointed on bimodal action of this compound, as responsible for uncoupling of oxidative phosphorylation and inhibition of electron transfer. Roots of tomato seedling treated with CAN were characterized by lower oxygen consumption than control plants and the effect was dose-dependent. Decreased respiration rate was observed just after 24 h of the culture period. This finding suggests CAN impact on root mitochondria, thus growth restriction could be due to low ATP synthesis. All the more, human cell lines exposed to CAN had reduced level of mitochondrial proteins and incorporation of CAN into mitochondrial proteins was linked to reduction of their stability (Konovalova et al., 2015). Application of CAN into growing medium of baby hamster kidney cells resulted in decrease of protein level and total DNA content (Wallace and Keir, 1986). RNA content was only slightly reduced. Our results indicated elevated level of total RNA especially observed after treatment with CAN at high (50 mM) concentration. Total soluble proteins level increased slightly after first 24 h of CAN treatment. Stress factors modulate mRNA translation. As was shown for roots of maize seedlings growing in anoxic conditions, mRNA was synthesized but its translation was limited (Fennoy et al., 1998). So, our data may imply CAN inhibitory effect on translation efficiency. Root growth and movement is under control of auxin, mostly IAA. Concentration of this hormone, and its polar transport implicate appropriate structure and function of roots. High cellular concentration of IAA results in restriction of root elongation. IAA application reversed inhibitory effect of CAN on adventitious root formation in isolated segments of hypocotyls of leafy spurge (Euphorbia esula L.) (Davis, 1997). Recently it has been shown that harmful effect of cyanamide on growth of roots of various plant species (tomato and corn) was related to the modifications in cell division and disturbances in IAA-ethylene balance (Soltys et al., 2011, 2012). Significant (threefold higher compared to the control) enlargement in IAA level was observed as fast reaction of tomato roots to cyanamide. On the contrary in Arabidopsis plants, citral led to increase in IAA level above 150% of the control after 10 ~ a et al., 2013). Our data indicate that CAN days of treatment (Gran altered IAA concentration just after 24 h of imbibition, and a high IAA level in CAN treated roots was kept up to 72 h. Thus, it suggests that besides influencing IAA metabolism CAN probably has also a strong impact on its polar transport. Especially that influence of coumarin on IAA polar transport was revealed (Lupini et al., 2014). Acropetal IAA transport via PINs (mostly PIN1) is disrupted also by elevated NO content, due to decline abundance of this proteins ndez-Marcos et al., 2011). (Ferna 5. Conclusion We provided the evidence that enhancement in ROS production accompanied by decline in RNS emission act as putative messengers in CAN induced reaction triggered by accumulation of auxins and resulting in disruption in root growth. Most of these effects (except RNS emission) may be identified rather as secondary modes of action of CAN. According to our data demonstrating high viability of cells exposed to CAN, we may assume that this NPAA may be used as a powerful modifier of NO level, application of which results in severe modification in cell physiology.
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Authors contributions The work was carried out in collaboration between all authors: AG and UK designed the experiments, coordinated all laboratory analysis, interpreted the results and wrote the manuscript. OA, PS, UK performed analysis, WB has done microscopic observation. All authors took part in interpretation of results and revised critically the manuscript. Acknowledgments Authors would like to thank R. Bogatek for fruitful suggestions during manuscript preparation, A. Antosik and M. Moczydłowski for their skilful technical assistance and B. Godley for English revision of the text. The study was funded by National Science Centre project no. 2014/13/B/NZ9/02074 and Ministry of Science and Higher Education Poland project no. DI2013012843. References Abd El-Gawad, H.M., Khalifa, A.E., 2001. Quercetin, coenzyme Q10, and L-canavanine as protective agents against lipid peroxidation and nitric oxide generation in endotoxin-induced shock in rat brain. Pharmacol. Res. 43, 257e263. http:// dx.doi.org/10.1006/phrs.2000.0781. Abrahim, D., Francischini, A.C., Pergo, E.M., Kelmer-Bracht, A.M., Ishii-Iwamoto, E.L., 2003. Effects of a-pinene on the mitochondrial respiration of maize seedlings. Plant Physiol. Biochem. 41, 985e991. http://dx.doi.org/10.1016/ j.plaphy.2003.07.003. Chomczynski, P., Sacchi, N., 1987. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162, 156e159. http://dx.doi.org/10.1006/abio.1987.9999. Colling, J., Stander, M.A., Makunga, N.P., 2010. Nitrogen supply and abiotic stress influence canavanine synthesis and the productivity of in vitro regenerated Sutherlandia frutescens microshoots. J. Plant Physiol. 167, 1521e1524. http:// dx.doi.org/10.1016/j.jplph.2010.05.018. Corpas, F.J., Barroso, J.B., 2015. Functions of nitric oxide (NO) in roots during development and under adverse stress conditions. Plants 4, 240e252. http:// dx.doi.org/10.3390/plants4020240. Corpas, F.J., Palma, J.M., del Río, L.A., Barroso, J.B., 2009. Evidence supporting the existence of L-arginine-dependent nitric oxide synthase activity in plants. New Phytol. 184, 9e14. http://dx.doi.org/10.1111/j.1469-8137.2009.02989.x. n, G., Pinedo, M., Di Rienzo, J., Novo-Uzal, E., Pomar, F., Lamattina, L., de la Corti Monzo Canal, L., 2014. Nitric oxide is required for determining root architecture and lignin composition in sunflower. Supporting evidence from microarray analyses. Nitric Oxide-Biol. Chem. 39, 20e28. http://dx.doi.org/10.1016/j.niox.2014.04.004. Davis, D.G., 1997. Polyamines, auxins and organogenesis in leafy spurge (Euphorbia esula L.). J. Plant Physiol. 151, 603e609. Ekanayake, S., Skog, K., Asp, N.-G., 2007. Canavanine content in sword beans (Canavalia gladiata): analysis and effect of processing. Food Chem. Toxicol. 45, 797e803. http://dx.doi.org/10.1016/j.fct.2006.10.030. Elsorra, E., Idris, E.E., Bochow, H., Ross, H., Borriss, R., 2004. Use of Bacillus subtilis as biocontrol agent. VI. Phytohormone like action of culture filtrates prepared from plant growth promoting Bacillus amyloliquefaciens FZB24, FZB42, FZB45 and Bacillus subtilis FZB37. J. Plant Dis. Prot. 111, 583e597. Fennoy, S.L., Nong, T., Bailey-Serres, J., 1998. Transcriptional and posttranscriptional processes regulate gene expression in oxygen-deprived roots of maize. Plant J. 15, 727e735. http://dx.doi.org/10.1046/j.1365-313X.1998.00249.x. ndez-Marcos, M., Sanz, L., Lewis, D.R., Muday, G.K., Lorenzo, O., 2011. Nitric Ferna oxide causes root apical meristem defects and growth inhibition while reducing PIN-FORMED 1 (PIN1)-dependent acropetal auxin transport. Proc. Nat. Acad. Sci. USA 108, 18506e18511. http://dx.doi.org/10.1073/pnas.1108644108. Gniazdowska, A., Krasuska, U., Andrzejczak, O., Soltys, D., 2015. Allelopathic compounds as oxidative stress agents: YES or NO. In: Gupta, K.J., Igamberdiev, A.U. (Eds.), Reactive Oxygen and Nitrogen Species Signaling and Communication in Plants, vol. 23. Springer, Verlag, Germany, pp. 155e176. http://dx.doi.org/ 10.1007/978-3-319-10079-1_8. ~ a, E., Sotelo, T., Díaz-Tielas, C., Krasuska, U., Bogatek, R., Reigosa, M.J., Sa nchezGran Moreiras, A., 2013. Citral induces auxin and ethylene-mediated malformations and arrests cell division in Arabidopsis thaliana roots. J. Chem. Ecol. 39, 271e282. http://dx.doi.org/10.1007/s10886-013-0250-y. Guan, Y., Lin, H., Ma, L., Yang, Y., Hu, X., 2014. Nitric oxide and hydrogen peroxide are important signals mediating the allelopathic response of Arabidopsis to p-hydroxybenzoic acid. Physiol. Plant 152, 275e285. http://dx.doi.org/10.1111/ppl.12164. Gupta, K.J., Igamberdiev, A.U., 2015. Compartmentalization of reactive oxygen
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