To gain more insight into the physiological function of nitrogen dioxide (NO2), we investigated the effects of exogenous NO2 on growth in Arabidopsis thaliana.
Plants were grown in air without NO2 for 1 wk after sowing and then grown for 1–4 wk in air with (designated treated plants) or without (control plants) NO2. Plants were irrigated semiweekly with a nutrient solution containing 19.7 mM nitrate and 10.3 mM ammonium.
Five-week-old plants treated with 50 ppb NO2 showed a ≤ 2.8-fold increase in biomass relative to controls. Treated plants also showed early flowering. The magnitude of the effects of NO2 on leaf expansion, cell proliferation and enlargement was greater in developing than in maturing leaves. Leaf areas were 1.3–8.4 times larger on treated plants than corresponding leaves on control plants. The NO2-induced increase in leaf size was largely attributable to cell proliferation in developing leaves, but was attributable to both cell proliferation and enlargement in maturing leaves. The expression of different sets of genes for cell proliferation and/or enlargement was induced by NO2, but depended on the leaf developmental stage.
Collectively, these results indicated that NO2 regulates organ growth by controlling cell proliferation and enlargement.
Nitric oxide (NO) and nitrogen dioxide (NO2) are often considered toxic gaseous air pollutants (Wellburn, 1990). For example, upon combustion of fuels, nitrogen (N) in the air is oxidized into NO, which is then rapidly converted into NO2 (Wellburn, 1990; Wallington & Nielsen, 1998). Atmospheric NO and NO2 have long been known to be either detrimental or beneficial to plants depending on the concentration and plant species (Capron & Mansfield, 1977; Sandhu & Gupta, 1989; Wellburn, 1990; Saxe, 1994). However, the molecular mechanisms of action have remained enigmatic, despite the fact that the rapid interconversion of these two N species in air suggests a similarity in their action (Wellburn, 1990).
Plants themselves also produce and emit NO2 (Klepper, 1979; Dean & Harper, 1986), and this may be an enzymatic or nonenzymatic process (Klepper, 1990). However, the physiological relevance of NO2 production has been largely unexplored apart from those pioneering studies (Leshem & Haramaty, 1996).
Recent research has established that NO is a phytohormone that influences various physiological processes. This has provided an answer to some long-sought questions regarding the activity of NO (Grün et al., 2006; Santner & Estelle, 2009). However, these developments have raised new questions about the role of NO2. Is the NO2 formed in plants simply an end product of no physiological relevance? Does NO2 play a physiological role similar to or different from that of NO? These questions are important not only for NO2 biology but also for the elucidation of the mechanisms and functions of NO.
Exogenously applied NO2 is known to regulate plant growth and development (Sandhu & Gupta, 1989; Takahashi et al., 2005, 2011). Prolonged exposure to exogenous NO2 at ambient concentrations can nearly double the total leaf area, nutrient uptake and shoot biomass in plants fed root N (Takahashi et al., 2005). Similar results have been reported in various plant species, including Arabidopsis thaliana (Arabidopsis) and various horticultural species (Ma et al., 2007; Adam et al., 2008; Takahashi et al., 2008; Xu et al., 2010). In addition, exogenous NO2 accelerates flowering time and increases flower number and fruit yield in tomato (Takahashi et al., 2011). However, N analyses of gaseous 15NO2-fed plants have indicated that the contribution of NO2 to total plant N is very minor (> 3%) and that NO2 may instead function as a signal rather than a nutrient (Takahashi et al., 2005).
In the present study, to gain more insight into the physiological function of NO2, we investigated the physiological, cellular and molecular mechanisms of growth regulation by exogenous NO2 in Arabidopsis. We first determined the optimal concentration of NO2, and quantified the effects of NO2 on shoot biomass and organ size. The growth of plant organs has been classified into two processes: cell proliferation (increase in cell number) and cell enlargement (increase in cell size; Potters et al., 2007; Granier & Tardieu, 2009; Kawade et al., 2010). Therefore, we analyzed which (either or both) process is responsible for NO2-induced increases in organ size and biomass. Furthermore, 23 genes for cell proliferation and/or cell enlargement were chosen, and their expression in response to NO2 was analyzed. In addition, the involvement of endoreduplication, a mechanism by which plants achieve increases in cell size (Melaragno et al., 1993; Sugimoto-Shirasu et al., 2005), in NO2-induced organ size control was also investigated.
We found that shoot biomass more than doubled, and leaf size increased 1.3- to 8.4-fold following NO2 treatment; however, the effects on leaf size, cell size and cell number depended on the leaf developmental stage. The NO2-induced increase in leaf size was attributable to increases in both cell number and size, and largely to cell size in maturing and developing leaves. Endoreduplication was not correlated with the NO2-induced increases in cell size. Quantitative real-time PCR analysis showed that the NO2-induced expression of different sets of genes for cell proliferation and/or cell enlargement depended on the leaf developmental stage. Collectively, our results indicated that NO2 regulates organ growth and size by controlling cell proliferation and enlargement.
Materials and Methods
We used a glass-walled NO2-exposure chamber (1.3 × 1.0 × 0.65 m in width, height and depth, respectively) model NOX-1000-SCII from Nippon Medical & Chemical Instruments Co. (Osaka, Japan) inside a growth room. Temperature, CO2 concentration and relative humidity in the chamber were set at 22 ± 0.1°C, 360 ± 30 ppm and 70 ± 1.5%, respectively. The air entering the chamber (at 1 l min−1) from outside was scrubbed of NO, NO2, and O3 (to 0 ppb) using activated charcoal and NaMnO4 (PureliteE30; Nippon Puretec Co., Tokyo, Japan).
Plant material and growth
Seeds of Arabidopsis thaliana (L.) Heynh. accession C24, or Columbia (Col-0), were surface-sterilized with 1.0% sodium hypochlorite, rinsed in pure water (18.0 MΩ), and sown in a rectangular plastic tray (22 × 5 × 20 cm in width, height and depth, respectively) containing vermiculite and perlite (1 : 1, v/v) and held in the NO2-exposure chamber described earlier. Seeds were allowed to germinate and grow for 1 wk in air without NO2. The tray was placed under fluorescent light (40 μmol photons m−2 s−1) with a 16 : 8 h, light : dark cycle. After the first week, we added NO2 (at concentrations of 0, 10 ± 0.2, 50 ± 0.3, 100 ± 20 or 200 ± 50 ppb) or 15NO2 (51.6 atom% 15N) at 50 ± 10 ppb to the air entering the exposure chamber. Seedlings were irrigated twice a week with half-strength inorganic salts of Murashige and Skoog medium (Murashige & Skoog, 1962) and grown for 1–4 wk more in the chamber.
Total carbon (C), N, phosphorus (P), potassium (K), calcium (Ca), magnesium (Mg), and sulfur (S) in plant samples were determined as described previously (Takahashi et al., 2005). Nitrogen analysis for 15NO2-fed plants was performed as described previously (Morikawa et al., 2004; Takahashi et al., 2005).
Leaves were fixed in formalin–acetic acid–alcohol (FAA) and cleared using a chloral hydrate solution (chloral hydrate, 200 g; glycerol, 20 g; H2O, 50 ml) following Tsuge et al. (1996). Whole leaves and leaf cells were observed under a stereoscopic microscope (MZ FLIII; Leica Microsystems, Wetzlar, Germany) and a Nomarski differential interference contrast compound microscope (ECLIPSE 80i; Nikon, Tokyo, Japan), respectively, as described in Horiguchi et al. (2005). Microphotographs were captured, and leaf area, cell density, and cell size were determined using ImageJ software (http://rsb.info.nih.gov/ij/). The density of palisade cells per unit area in the subepidermal layer in the center of the leaf blade between the midvein and the leaf margin was determined for each leaf. This value was multiplied by the leaf area to calculate the total number of palisade cells. This determination was repeated on five individuals. To estimate cell size, 20 palisade cells per leaf (in the same region of the blade) were measured on five individuals. Transverse sections were prepared following Tsuge et al. (1996). Leaf samples were fixed in FAA and dehydrated using a 50, 60, 70, 80, 90, 99.5 and 100% v/v ethanol series, followed by incubation in Technovit 7100 resin : ethanol (1 : 1, v/v; Heraeus Kulzer, Tokyo, Japan) at room temperature for c. 1 h, then kept at 25°C overnight in 100% Technovit 7100 resin to ensure resin penetration. Hardening of resin and embedding in 100% Technovit 7100 resin were performed following instructions from the Technovit 7100 resin supplier. Sections of 5–10 μm thickness were cut with a rotary microtome (RM2245; Leica Microsystems), and stained with 0.1% (w/v) Toluidine blue in 50 mM sodium phosphate (pH 7.0) for 1 min, then rinsed with tap water. After drying, the sections were mounted in Entellan Neu (Merck, Darmstadt, Germany) and photographed under bright-field illumination using a Nikon ECLIPSE 80i microscope.
Flow cytometric analysis
We performed flow cytometric analysis following Kozuka et al. (2005). The blade of the eighth leaf on each 5-wk-old plant that had been frozen and stored at −80°C was chopped with a razor blade in a plastic Petri dish containing 500 μl cold buffer (10 mM Tris–HCl, pH 8.0, 2 mM MgCl2, 0.1% Triton X-100, and 3% (w/v) polyvinylpyrrolidone (PVP), 1% (v/v) 2-mercapto-ethanol) and incubated on ice for 20 min. The suspension was filtered through a 20 μm nylon mesh (CellTrics; Partec, Münster, Germany), treated with RNase A (100 μg ml−1) at 37°C for 30 min, and stained with propidium iodine (40 μg ml−1) at 4°C for 1 h. The nuclear suspension was then subjected to flow cytometry with an Epics XL flow cytometer (Beckman Coulter, Fullerton, CA, USA) following the manufacturer's instructions. Three independent measurements were performed for each sample.
Gene expression analysis
Blades of Arabidopsis rosette leaves frozen in liquid nitrogen and stored at −80°C were homogenized and used for total RNA extraction with an RNeasy Plant Mini Kit (Qiagen) following the manufacturer's instructions. The residual DNA was removed by on-column DNase digestion with RNase-Free DNase (TaKaRa, Otsu, Shiga, Japan). One microgram of total RNA was reverse-transcribed with the PrimeScript RT Reagent Kit for Real Time (TaKaRa) in a 10 μl reaction volume. Quantitative real-time PCR was performed with Power SYBR Green PCR Master Mix and an ABI 7300 real-time PCR system (Applied Biosystems, Foster City, CA, USA) following the manufacturer's protocol. The RT reaction mixture was diluted 50-fold with pure water, and 4 μl was added (as a template for amplification) to a 20 μl total reaction volume containing 10 μl SYBR Green mix and 0.3 μM each specific primer. Cycling parameters were as follows: initial denaturation at 95°C for 15 min, followed by 40 cycles of 15 s each at 95°C, and 1 min at 60°C. The specificity of the PCR was confirmed by melting-curve analysis. Relative gene expression was calculated using the comparative method and UBQ10 as a reference gene. The list of primer pairs used is provided in Supporting Information, Table S1.
GraphPad Prism 6.0 (GraphPad Software, La Jolla, CA, USA) was used for all statistical analyses. Student's t-test or Mann–Whitney U-test was used to compare two groups. Dunnett's t-test was used to compare three or more groups against a single control. Pearson's correlation analysis with Bonferroni correction was performed between log-transformed data of the leaf area, cell size and cell number. P <0.05 was considered to indicate statistical significance.
Increases in shoot biomass and acceleration of flowering upon exposure to NO2
Plants were always grown in the absence of NO2 for the first week after sowing, and then grown for 1–4 wk in the presence (designated +NO2-treated plants) or absence (−NO2 control plants) of NO2. Plant age is given here in wk after sowing and refers to the time of harvest. Accession C24 was used unless otherwise stated.
We first determined the effect of NO2 concentration on the yield of shoot biomass in 4-wk-old plants (Table 1). Shoot biomass of +NO2-treated plants under 10 ± 0.2 and 50 ± 0.3 ppb NO2 was c. 2.5-fold higher than that of −NO2 control plants. This enhancement effect was greater than the value (1.1 times) reported previously by Xu et al. (2010). Treatments of 100 ± 20 and 200 ± 50 ppb NO2 produced no stimulating effects or marginally inhibited the growth of plants (Table 1). Hence, we used 50 ± 0.3 ppb NO2 treatments in subsequent experiments.
Table 1. Effect of nitrogen dioxide (NO2) concentration on the yield of shoot biomass of 4-wk-old plants of Arabidopsis thaliana accession C24
NO2 concentration (ppb)
DW of shoots (mg per plant)
After sowing, plants were grown for 1 wk in air not containing NO2, and then grown for a further 3 wk in air containing 0, 10 ± 0.2, 50 ± 0.3, 100 ± 20 or 200 ± 50 ppb NO2. Data are means ± SD. Statistical significance assessed by Dunnett's t-test: ***, P <0.001.
We determined the time courses of shoot biomass accumulation in +NO2 and −NO2 control plants during 1–4 wk of exposure (Fig. 1). At week 1, shoot biomass did not significantly differ between treatments, indicating that 1 wk of plant exposure is too short to produce an effect. From week 2 onward, the shoot biomass of +NO2-treated plants became significantly greater than that of −NO2 control plants. By week 3, a 2.2-fold difference in shoot biomass was observed between +NO2 and −NO2 treatments, an outcome very similar to the experiment reported in Table 1. The difference in shoot biomass between +NO2 and −NO2 treatments increased markedly by the fourth week of exposure; by then, the difference had reached 2.8-fold (see Fig. 1). Images of typical 5-wk-old +NO2 and −NO2 control plants are presented in Fig. 2. NO2 treatment also increased root biomass by 1.9-fold; the root DWs were 4.8 ± 0.8 and 2.5 ± 0.6 mg per plant (mean of 10 plants ± SD, P <0.001) in 5-wk-old +NO2-treated plants and −NO2 control plants, respectively.
Shoot biomass of Arabidopsis accession Col-0 also increased upon NO2 treatment (Fig. S1). This is in agreement with a previous report by Xu et al. (2010). Treated 4-wk-old plants of Col-0, which were close to the end of vegetative growth, had shoot biomasses 1.7-fold higher than controls: 24.2 ± 5.5 and 14.3 ± 2.5 mg (mean ± SD, n =5) for +NO2-treated plants and −NO2 control plants, respectively. The different responses to NO2 treatment between Col-0 and C24 are applicable to genetic studies in Arabidopsis.
Flowering time was significantly shortened after NO2 treatment in both C24 and Col-0 accessions (Fig. 3). The number of days after sowing when flower bolts were 1 cm in length was used as a measure of flowering time (Kotchoni et al., 2009). Flowering time of C24, a late flowering accession, had median values of 41 and 42 d in +NO2-treated plants and −NO2 control plants, respectively. Student's t-test of the respective mean ± SD values (40 ± 3 and 42 ± 3 d; n =20) showed that this difference was statistically significant (P <0.05) (Fig. 3). However, the effect of NO2 treatment was more distinct on flowering time of Col-0, an early flowering accession. Median Col-0 flowering times were 34 and 40 d in +NO2-treated plants and −NO2 control plants, respectively. Student's t-test of the respective mean ± SD values (34 ± 1 and 40 ± 3 d; n =17–26) showed that this difference was statistically significant (P <0.001) (Fig. 3). Similar flowering acceleration has been reported in tomato: NO2 accelerates flowering time, and increases flower number and fruit yield (Takahashi et al., 2011). These results of early flowering by NO2 treatment run counter to the reported results of late flowering induced by NO (He et al., 2004; see discussion later).
Stimulation of shoot growth by NO2 treatment was accompanied by increased shoot uptake of major nutrients, such as C, N, P, K, Ca, Mg and S. The contents of these nutrients per DW of shoot were almost equivalent for +NO2-treated plants and −NO2 control plants, and on a per-shoot basis their contents almost doubled (Table S2). These results are congruent with our previous work in Nicotiana plumbaginifolia (Takahashi et al., 2005) and other horticultural species (Adam et al., 2008).
The content of N derived from NO2 (NO2–N) in above-ground parts of plants was determined after feeding with 15N-labeled gaseous NO2 and unlabeled nitrate. The 15N : 14N ratio gives a measure of the content of NO2–N in total plant N. Using mass spectrometry (Morikawa et al., 2004), we determined that NO2–N comprised < 5% (4.05 ± 0.75%; mean ± SD, n =3) of the total N in the shoots of +NO2-treated plants. Therefore, the role of NO2–N as an N source was clearly minor, indicating that NO2 may function instead as an environmental signal to stimulate the growth of plants. This is in line with our previous reports (Takahashi et al., 2005; Adam et al., 2008).
Increase in leaf size in response to NO2
The increased total leaf area in response to NO2 treatment (Table S2) indicated that leaf size must have increased after NO2 treatment. Therefore, we analyzed the sizes of rosette leaves in positions 1–25 on 5-wk-old +NO2-treated plants and −NO2 control plants, which had 28 and 25 rosette leaves, respectively. In both treatments and controls, leaves 1–11 appeared to be close to maturity, while leaves 12–25 were in the developing stages. We measured the areas of leaf blades (Fig. 4a). Numerical details of leaf area determinations are provided in Table S3.
The distribution of leaf area against leaf position showed a right-skewed bell-shaped pattern on −NO2 control plants (Fig. 4a, open columns), while this skewness was less pronounced and the distribution became less asymmetric for +NO2-treated plants (Fig. 4a, closed columns). The location of the mode of distribution shifted to the younger leaf side by about four units (from position 8 to position 12) after NO2 treatment, which suggested that the NO2 effect on leaf area was more distinct in younger leaves than in older leaves (see later).
The areas of leaves 1 (the oldest) to 25 (the youngest) were greater (1.3–8.4 times) in +NO2-treated plants than in −NO2 control plants in the corresponding leaf positions (Fig. 4a). These differences were statistically significant at all positions (Fig. 4a). Therefore, the NO2 treatment significantly increased leaf sizes in both developing (positions 12–25) and maturing (positions 1–11) leaves.
The NO2-induced fold-change in leaf area (FCLA) was estimated by dividing the leaf area of +NO2-treated plants by that of −NO2 control plants from the corresponding leaf position. The FCLA varied from 1.3 to 2.5 in leaves 1–11, while a larger FCLA of 2.9–8.4 was observed in leaves 12–25 (Table S3). Mann-Whitney U-test indicated that the difference in the mean FCLA between leaves 1–11 and 12–25 was significant (P <0.01). This result indicated that the magnitude of the NO2 effect on leaf expansion changed as a function of the developmental stage, and was more distinct in developing leaves than in maturing leaves.
Increases in leaf cell number and size in response to NO2
The size of an organ is determined by the number and size of its constituent cells (Potters et al., 2007; Granier & Tardieu, 2009; Kawade et al., 2010). Therefore, we determined whether the NO2-induced increase in leaf area was attributable to increases in cell number or cell size, or both. To this end, we analyzed the palisade cells in the adaxial subepidermal layer, as, in Arabidopsis, this layer consists of cells neatly aligned in the paradermal plane throughout leaf development (Tsuge et al., 1996). This was the case in both +NO2-treated and −NO2 control plants, as shown in Fig. 5. Thus, we determined the sizes and numbers of palisade cells in leaves occupying positions 1–25 in 5-wk-old +NO2-treated plants and −NO2 control plants. The results are shown in Fig. 4(b,c). Numerical details for cell size and number determinations are given in Table S3.
In contrast to the distribution of leaf area, the distribution of cell size was not bell-shaped, but decreased monotonically with leaf position in both treated and control plants (Fig. 4b), indicating that cell size increased almost linearly with age. Also, NO2 treatment did not affect this trend (see Fig. 4b). The differences in cell size were statistically significant at all positions, other than leaf 2 (Fig. 4b). Therefore, NO2 treatment significantly increased the cell sizes in both developing and maturing leaves.
The NO2-induced fold-change in cell size (FCCS) was estimated by dividing the cell size of +NO2-treated plants by that of −NO2 control plants at the corresponding leaf position (Table S3). The FCCS (2.0–3.2) of leaves 12–25 appeared to be larger than that (1.3–1.9) of leaves 1–11. The Mann–Whitney U-test indicated that the difference in the mean FCCS between leaves 12–25 and 1–11 was significant (P <0.01). This result indicated that the magnitude of the NO2 effect on cell enlargement changed as a function of developmental stage, and was more distinct in developing leaves than in maturing leaves. This is in agreement with the greater effect of NO2 on leaf expansion in younger leaves than older leaves.
The distribution of cell number against leaf position was more or less bell-shaped for both +NO2-treated plants and −NO2 control plants (Fig. 4c), which was similar to that of leaf size, but different from that of cell size. In contrast to leaf area or cell size, differences in cell numbers between treatment and control plants were not significant in almost all older leaves (positions 1–11 other than leaf 4), suggesting that the final cell numbers in leaves were not affected by NO2 treatment. Differences in cell number were significant in almost all younger leaves (positions 12–25 other than leaves 12 and 25), indicating that NO2 treatment significantly increased cell numbers in developing leaves.
The NO2-induced fold-change in cell number (FCCN) was estimated by dividing the cell number of +NO2-treated plants by that of −NO2 control plants at the corresponding leaf position (Table S3). The FCCN (1.2–3.1) of leaves 12–25 was larger than that (0.9–1.5) of leaves 1–11. The Mann–Whitney U-test indicated that the difference in the mean FCCN between leaves 12–25 and 1–11 was significant (P <0.01). This result indicated that the magnitude of the NO2 effect on cell proliferation changed as a function of developmental stage, and was more distinct in developing leaves than maturing leaves.
Therefore, it is clear that the NO2 treatment induced leaf expansion, cell proliferation and enlargement in a developmental stage-dependent manner, and that the magnitudes of the NO2 effect on leaf expansion, cell proliferation and enlargement were greater in developing leaves than maturing leaves.
Are increases in leaf area attributable to increases in cell size and/or number?
To explore whether increases in leaf area following NO2 treatment were attributable to increases in cell sizes and/or numbers, correlations between FCLA and FCCS (or FCCN) were analyzed. The FCLA, FCCS and FCCN values were transformed into logarithmic values, which were subjected to Pearson's correlation analysis with Bonferroni correction for multiple comparisons (Table 2).
Table 2. Correlation between log10 FCLAa and log10 FCCSb (or log10 FCCNc) in leaves 1–25, leaves 1–11 and leaves 12–25 of Arabidopsis thaliana accession C24
P after Bonferroni correction
FCLA (fold-change in leaf area) = (leaf area of +NO2-treated plants)/(leaf area of −NO2 control plants).
FCCS (fold-change in cell size) = (cell size of +NO2-treated plants)/(cell size of −NO2 control plants).
FCCN (fold-change in cell number) = (cell number of +NO2-treated plants)/(cell number of −NO2 plants).
The correlation between log FCLA and log FCCS was high and significant in leaves 1–25 (R =0.9, P <0.001). When this correlation was evaluated separately in younger leaves (12–25) and older leaves (1–11), it was found that the correlation between NO2-induced increases in leaf areas and cell enlargement was stronger in older leaves (R =0.7, P <0.05) than in younger leaves (R =0.3, P >0.5) (Table 2). These results indicated that NO2-induced increases in leaf areas were correlated with cell enlargement, and that the correlation was stronger in maturing leaves than in developing leaves.
The correlation between log FCLA and log FCCN was also high and significant in leaves 1–25 (R =0.9, P <0.001). Similar correlation results were obtained in both younger (12–25) (R =0.9, P <0.001) and older (1–11) (R =0.7, P <0.05) leaves (Table 2). These results indicated that NO2-induced increases in the areas of both developing and maturing leaves were correlated with cell proliferation.
Based on these results, NO2-induced increases in leaf areas were largely attributable to cell proliferation in developing leaves, while the effect was attributable to both cell proliferation and enlargement in maturing leaves.
This conclusion is in a good agreement with the proposed mechanism for the formation of larger leaves (Gonzalez et al., 2012; Powell & Lenhard, 2012). In this mechanism, the increase in leaf size in the early stages of leaf development is mainly attributable to cell proliferation (including increased cell division rate and prolonged proliferation). The increase in leaf size during the later stages of leaf development was attributable to cell proliferation and enlargement-related events, such as an increased expansion rate and prolonged expansion (Gonzalez et al., 2012; Powell & Lenhard, 2012).
Contribution of endoreduplication to cell enlargement
A mechanism used by plants to achieve increases in cell size is to increase the ploidy level through successive rounds of DNA replication without cell division, a process called endoreduplication (Melaragno et al., 1993; Sugimoto-Shirasu et al., 2005). To explore the involvement of endoreduplication in NO2-induced cell enlargement, we determined the ploidy levels in +NO2-treated and −NO2 control plants using leaves in position 8 on 5-wk-old plants as sample material for flow cytometry (Fig. 6).
Five peaks of fluorescence corresponding to 2C, 4C, 8C, 16C and 32C DNA contents were observed in the both +NO2-treated and −NO2 control plants. The distributions of the ploidy levels were very similar in treated and control plants (Fig. 6). The most frequent nuclei (33.0 ± 4.7 and 39.7 ± 5.3%, respectively) in both controls and treatment subjects had 8C DNA content, and the proportions of nuclei with the remaining four ploidy levels were very similar between experimental groups (Fig. 6). A very similar outcome was obtained in the analysis of the endoreduplication factor, which gave a measure of the mean number of endocycles per 100 cells, that is, 163.5 ± 9.1 and 168.2 ± 4.2 (n =3) for −NO2 control and +NO2-treated plants, respectively. Therefore, NO2-induced cell enlargement did not correlate with endoreduplication.
Expression of cell proliferation and/or enlargement genes in response to NO2
Several genes involved in increased organ size and biomass upon ectopic overexpression (for positive regulators) or down-regulation (for negative regulators) have been investigated using Arabidopsis as a model plant (reviewed by Gonzalez et al., 2009, 2012; Powell & Lenhard, 2012). Investigations using overexpressors of positive regulators or down-regulated transgenic plants of negative regulators have shown that multiple pathways independently converge on organ size control in Arabidopsis (Gonzalez et al., 2009, 2012; Powell & Lenhard, 2012).
To determine which genes are involved in NO2-induced leaf size increase, we focused on nine (ABAP1, ANT, AN3/AtGIF1, ARGOS, AVP1, DA1, GRF5, KLU and UBP15), 11 (ARL, ATAF2, ATHB16, EXO, EXP10, GRF1, GRF2, NAC1, OBP2, RON2 and TOR), and three (ARF2, EBP1 and GA20OX1) genes that stimulate cell proliferation, enlargement, and both cell proliferation and enlargement, respectively (Gonzalez et al., 2009, 2012), and analyzed (by quantitative real-time PCR) their average transcript expression levels in young (leaves 21–25), middle (leaves 12–20), and old (leaves 1–11) leaves of 5-wk-old +NO2-treated plants and −NO2 control plants (Fig. 7).
In young leaves, three positive regulators for cell proliferation (AVP1; Li et al., 2005), enlargement (NAC1; Xie et al., 2000), and both cell proliferation and enlargement (GA20OX1; Huang et al., 1998) were significantly up-regulated in +NO2-treated plants compared with the controls (Fig. 7).
In middle leaves, a negative regulator of cell proliferation (DA1; Li et al., 2005) was significantly down-regulated in +NO2-treated plants compared with the controls (Fig. 7).
In old leaves, a totally different set of genes was induced by NO2, including GA20OX (Fig. 7). Three positive regulators of cell proliferation (ARGOS (Hu et al., 2003), GRF5 (Horiguchi et al., 2005), and KLU (Anastasiou et al., 2007)) showed 100-fold increased gene expression in +NO2-treated plants compared with control plants. Furthermore, two positive regulators of cell proliferation (AN3/AtGIF1 (Horiguchi et al., 2005) and ABAP1 (Masuda et al., 2008)) showed less dramatic but distinct (two- to fivefold higher) gene expression in treated plants than in control plants. Moreover, three positive regulators of cell enlargement (ARL (Hu et al., 2006), EXO (Schröder et al., 2009), and EXP10 (Cho & Cosgrove, 2000)) and GA20OX showed 20- to 50-fold higher gene expression in +NO2-treated plants than in control plants. In addition, negative regulators of enlargement (RON2; Cnops et al., 2004) and of both proliferation and enlargement (ARF2, Schruff et al., 2006) showed smaller but distinct differences (two- to fivefold) in expression levels between +NO2-treated plants and control plants (Fig. 7).
Of the 23 genes investigated in the present study, nine showed a lack of either significant (ANT, UBP15, ATHB16, GRF1, GRF2 and EBP1) or physiologically meaningful (OBP2, ATAF2 and TOR) differences in transcript expression levels between +NO2-treated plants and control plants. The genes OBP2 (Skirycz et al., 2006), ATAF2 (Delessert et al., 2005), and TOR (Deprost et al., 2007) showed significantly augmented (OBP2) or diminished (ATAF2 and TOR) expression. However, because these genes are known to be negative (OBP2) or positive regulators (ATAF2 and TOR), they are not likely to be involved in the NO2-induced increases in organ size and biomass.
Clearly, all of the 14 genes that showed significant and physiologically meaningful differences in transcript expression levels between +NO2-treated plants and control plants were differentially regulated by the developmental stages of Arabidopsis leaves. The following genes were likely involved in NO2-induced organ size control: AVP1, GA20OX1, DA1 and NAC1 in developing leaves and ABAP1, AN3/AtGIF1, ARGOS, GRF5, KLU, ARL, EXO, EXP10 GA20OX, RON and ARF2 in maturing leaves.
NO2 is a signal that triggers plant growth and development
Exogenous NO2 influences various physiological processes in plants. Exposing plants that were well fed with soil N to gaseous NO2 increased the uptake of nutrients, photosynthesis, and nutrient metabolism so that shoot biomass, total leaf area, and shoot contents per shoot of C, N, P, K, Ca, Mg and S approximately doubled in comparison to control plants (Table S2), as has been shown in previous studies (Takahashi et al., 2005; Adam et al., 2008). Increase in the photosynthetic rate by NO2 has also been reported elsewhere (Ma et al., 2007; Xu et al., 2010). Mass spectrometric analysis of the 15N : 14N ratio showed that NO2-derived N (NO2–N) comprised 3–5% of the total plant N, indicating that the contribution of NO2–N to total N was very minor, as reported in previous studies (Takahashi et al., 2005; Adam et al., 2008). Taken together with the results that NO2 induced increases in leaf size and biomass through differential enhancement of cell proliferation and enlargement, NO2 may function as a signaling molecule rather than an N source.
Similarities and differences between the effects of NO and NO2
A very similar increase in Arabidopsis shoot biomass was obtained upon plant exposure to NO gas at the same concentration as NO2 (Fig. S2). This is in agreement with previous studies reporting that treatment of Arabidopsis seedlings with an NO donor, sodium nitroprusside, enhances vegetative growth (He et al., 2004) and that exposure to NO gas promotes the expansion of pea leaf discs (Leshem & Haramaty, 1996) and the vegetative growth of spinach (Jin et al., 2009). Taken together, these results suggest that the effects of NO2 mimic those of NO by stimulating the vegetative growth of Arabidopsis, and that NO and NO2 share a very similar or identical mechanism for the stimulation of vegetative growth.
By contrast, NO and NO2 have opposite effects on flowering time. Exogenous NO delays the flowering of Arabidopsis Col-0 (He et al., 2004), whereas exogenous NO2 accelerates flowering time in Arabidopsis Col-0 and C24 (Fig. 3) and tomato (Takahashi et al., 2011). As it is known that C24 and Col-0 have different compositions of alleles of flowering time genes FRIGIDA (FRI) and FLOWERING LOCUS C (FLC) (Sanda & Amasino, 1995), the fact that the magnitude of the effect of NO2 on acceleration of flowering was more distinct in early flowering Col-0 than in late flowering C24 (Fig. 3) suggests that these genes may be involved in the NO2 effect of accelerating flowering in Arabidopsis.
The reciprocal effects of NO and NO2 on flowering time in Arabidopsis provide evidence that their interconversion inside and outside cells is limited (see later). GA stimulates vegetative growth and flowering (Mutöasa-Gottgens & Hedden, 2009), and plants overexpressing GA20OX1 exhibit stimulated vegetative growth and early flowering, which is attributable to increased concentrations of active GAs (Huang et al., 1998; Coles et al., 1999; Gonzalez et al., 2010, 2012). Therefore, our results of stimulated vegetative growth and early flowering in Arabidopsis can be attributed, at least in part, to the up-regulation of GA20OX1 in developing and maturing leaves in response to NO2 treatment (see Fig. 7).
In atmospheric chemistry, a rapid interconversion of NO and NO2 is known as the Leighton relationship, in which oxidation of NO by ozone (1.1 × 1010 M−1 s−1) and photolysis (1 × 10−2 s−1) of NO2 by sunlight play important roles. The lifetime of NO and NO2 are estimated to be 1.3 and 2–6 min, respectively (Wellburn, 1990; Wallington & Nielsen, 1998). In the absence of ozone or in ozone-depleted air, NO is autoxidized to NO2 by atmospheric oxygen at a much lower rate (three orders of magnitude lower than in the presence of ozone).
By contrast, the biological lifetime of NO is reportedly 7–27 s (Gupta et al., 2011). In biological systems, endogenous NO can be rapidly converted to NO2 via reaction with in vivo oxidants such as superoxide at a similar rate to NO and ozone. In addition, NO can also be autoxidized at a high rate (1.4 × 109 M−2 s−1) in the hydrophobic lipid phase of the membrane (Shapiro, 2005). Enzymatic and nonenzymatic processes that convert endogenous NO2 to NO have been identified (Purwantini & Mukhopadhyay, 2009). However, the extent to which such processes contribute quantitatively to endogenous NO concentrations remains unknown. Nonetheless, these facts imply a close relationship between the in vivo effects of NO and NO2, and indicates that understanding the effects of NO2 could increase our understanding of the effects of NO, and vice versa.
Importance of identification of the NO2 target site and biosynthesis pathway
It has been reported that NO induces numerous genes involved in various plant signaling pathways, and identifying the correct response evoked by NO is a major challenge (Grün et al., 2006). The identification of target molecule(s) of NO2 is crucial to identifying its specific mechanism of action. However, the sites of action, receptors, and signal transduction cascades of NO2 have not been identified. The reported fact that NO2 is involved in protein tyrosine nitration as a proximal intermediate in plants (Shapiro, 2005) suggests that nitrated proteins are a potential target of NO2 action. Direct evidence for the involvement of NO2 in intracellular redox signaling has been reported in humans and rodents. In the endoplasmic/sarcoplasmic reticulum, Ca2+–Mg2+-adenosine triphosphatase (SERCA2) acts as a sensor of NO2, and the tyrosine residues Tyr294 and Tyr295 are reversibly nitrated and denitrated in response to the cellular redox state (Bigelow, 2009). However, whether these mechanisms established in animals are true in plants has yet to be elucidated. Besides the involvement of NO2 in protein tyrosine nitration, its metabolism in plants has also been reported (Morikawa et al., 2004, 2005; Takahashi et al., 2005).
Although more than half a dozen NO production pathways or routes (Fröhlich & Durner, 2011) exist in plants, and many studies have reported on NO2 emission by plants (Klepper, 1979, 1990; Dean & Harper, 1986), in vivo biosynthetic pathways for NO2 are largely unexplored, other than reports on the enzymatic production of NO2 (Klepper, 1990). On the other hand, in vitro production of NO2 via one-electron oxidation of nitrite by hemeproteins such as horseradish peroxidase (Shibata et al., 1995) and leukocyte peroxidases (Brennan et al., 2002), Arabidopsis hemoglobins (Sakamoto et al., 2004), and human hemo/myoglobins (Turko & Murad, 2002) has been demonstrated. However, the extent to which these enzyme-mediated reactions contribute to the in vivo production of NO2 is not known.
In conclusion, determining whether NO2 plays a hormonal role requires further study, which should include the identification of target sites or receptors of NO2 and the in vivo biosynthetic pathways of NO2. However, based on the results of the present study, it is clear that NO2 regulates organ growth and size in Arabidopsis by controlling cell proliferation and enlargement.
This work was supported in part by a Grant-in-Aid for Scientific Research from the Japan Society for the Promotion of Science and by CREST, of the Japan Science and Technology Agency. We thank NIBB Core Research Facilities, National Institute for Basic Biology, Myodaiji, Okazaki, Japan, for use of the flow cytometer.