• We report the unexpected novel finding that exogenously supplied atmospheric NO2 at an ambient concentration is a plant vitalization signal to double shoot size and the contents of cell constituents.
• When seedlings of Nicotiana plumbaginifolia were grown for 10 wk under natural light and irrigation with 10 mm KNO3 in air containing (+NO2 plants) or not containing (–NO2 plants) 15NO2 (150 ± 50 ppb), shoot biomass, total leaf area, and contents per shoot of carbon (C), nitrogen (N), sulphur (S), phosphorus (P), potassium (K), calcium (Ca), magnesium (Mg), free amino acids and crude proteins were all approximately 2 times greater in +NO2 plants than in –NO2 plants.
• In mass spectrometric analysis of the 15N/14N ratio, it was found that NO2-derived N (NO2-N) comprised < 3% of total plant N, indicating that the contribution of NO2-N to total N was very minor.
• It thus seems very likely that the primary role of NO2 is as a multifunctional signal to stimulate plant growth, nutrient uptake and metabolism.
Atmospheric nitrogen dioxide (NO2) is regarded as an air pollutant (Wellburn et al., 1997), with most urban NO2 reported to originate from the exhaust gas of vehicles (Yunus et al., 1996). However, we recently reported (Morikawa et al., 2004) that about one-third of NO2-derived N (NO2-N) taken up into plants is converted into a previously unknown Kjeldahl-unrecoverable organic nitrogen (designated unidentified nitrogen or UN), and that the UN-bearing compounds include a novel heterocyclic Δ21,2,3 thiadiazoline derivative and nitroso- and nitro-organic compounds (Miyawaki et al., 2004; Morikawa et al., 2005). These results strongly suggest that exogenously supplied or atmospheric NO2 is not a simple supplemental N source or a pollutant, but instead functions as an airborne reactive nitrogen species signal (Morikawa et al., 2004, 2005). Consistent with this possibility are the reports that endogenously produced nitrogen oxides (NOx) such as nitric oxide (NO) function as a vital plant signal (Wendehenne et al., 2001; Neill et al., 2003; Sakamoto et al., 2004). However, whether exogenously supplied atmospheric NOx plays a role as a plant signal has never been addressed.
We have investigated plant-mediated mitigation of atmospheric NO2 pollution and the use of NO2 as an alternative N fertilizer (Morikawa et al., 1998; Goshima et al., 1999; Takahashi et al., 2001, 2003; Morikawa et al., 2004). In our attempt to produce NO2-philic plants that can grow with NO2 as the sole nitrogen source (Morikawa et al., 2003), we unexpectedly discovered that, although NO2-N taken up by plants during growth for 10 wk in air containing 150 ppb 15NO2, which corresponds to heavily polluted urban air (OECD, 2002), comprised only a minor part (< 3%) of total plant N, exogenously supplied NO2 almost doubled biomass and total leaf area, the contents of carbon (C), nitrogen (N), sulphur (S), phosphorus (P), potassium (K), calcium (Ca) and magnesium (Mg), and the contents of free amino acids and crude proteins.
Materials and Methods
Fumigation chamber and conditions
A glass-walled NOx-fumigation chamber (1.5 × 1 × 0.7 m in width, height and depth; model HM1500; Nippon Medical & Chemical Instruments Co., Osaka, Japan) was placed in a confined glasshouse (6.9 × 2.4 × 3.0 m in width, height and depth; model BTH-P1-TH; Nippon Medical & Chemical Instruments Co.). The air entering the glasshouse (at a rate of 4 m3 min−1) from outside was cleaned of NO2 and O3 by activated charcoal. NO2 and O3 concentrations in the glasshouse were < 5 ppb. Temperature, CO2 concentration and relative humidity were controlled at 22 ± 0.3°C, 340 ± 80 ppm and 70 ± 4%, respectively, by a climate controlling system (CTH-G 100; Nippon Medical & Chemical Instruments Co.). The air entering the fumigation chamber (at a rate of 1 l min−1) was mixed with 15N-labelled (51.6 atom% 15N) NO2 at a final concentration of 150 ± 50 ppb. Temperature, CO2 concentration and relative humidity in the chamber were controlled by an independent climate controlling system (CTH-C 70; Nippon Medical & Chemical Instruments Co.) at the same values as those for the glasshouse.
Plant material and growth
Seeds of Nicotiana plumbaginifolia Viviani were surface-sterilized with 2.5% sodium hypochlorite, rinsed with pure water (18.0 MΩ) and sown in Petri dishes containing B-medium (Vaucheret et al., 1992) solidified with 0.3% Gellangum (Wako Pure Chemical Ind., Osaka, Japan). After germination, seedlings were grown under continuous fluorescent light (70 µmol photons m−2 s−1) at 22 ± 0.3°C and a relative humidity of 70 ± 4% for 2 wk. Germination of seeds and aseptic culture of the seedlings took place in a growth chamber (model ER-20-A; Nippon Medical & Chemical Instruments Co.) as described previously (Morikawa et al., 2004).
The seedlings were then transferred to plastic pots containing vermiculite and perlite (1 : 1, volume/volume) and grown in the glasshouse (< 5 ppb NO2) under natural light for 1 wk, after which plants were divided into two groups (of six plants each). One group continued to grow in the glasshouse for a further 10 wk (designated –NO2 plants). The other group was placed inside the fumigation chamber (150 ± 50 ppb NO2) and allowed to grow under natural light for a further 10 wk (designated +NO2 plants). Both groups of plants were irrigated with Coïc & Lesaint (1971) medium containing 10 mm KNO3 every 4 d. Total leaf area (see next section) was measured weekly. Harvested plants were washed with pure water, separated into shoots and roots, lyophilized, powdered and stored in a desiccator until use, as reported elsewhere (Takahashi et al., 2001).
Determination of total leaf area
The total leaf area of each plant was estimated from the areas of individual leaves, calculated from leaf length (LL) and leaf width (LW) using the following equation (Hammer et al., 1998):
Individual leaf area = 0.8 × (LL × LW)
The factor of 0.8 was estimated from the comparison of calculated leaf area and directly measured leaf area (for about 40 leaf samples) using a leaf area meter (CI-203; CID Inc., Camas, WA, USA) with an attachment (CI-203CA).
Total N and C in plant samples were determined using an EA-MS analyzer consisting of an elemental analyser (EA1108 CHNS/O; Fisons Instruments, Milan, Italy) directly connected to a mass spectrometer (Delta C; Thermo-Finnigan, Bremen, Germany) as described elsewhere (Morikawa et al., 2004). After being wet-ashed in a HNO2:H2O2 mixture, as reported elsewhere (Rodushkin et al., 1999), plant samples were analysed for total P, K, Ca and Mg by inductively coupled plasma-atomic emission spectroscopy (ICP-AES; model 8100; Shimadzu, Kyoto, Japan). For the analysis of total S, plant samples were combusted in an oxygen combustion bomb, and SO2 released was recovered in 0.3% H2O2 and analysed by ICP-AES. Total free amino acids were analysed as reported previously (Igarashi et al., 2003). Crude protein content was analysed by the Kjeldahl method as reported elsewhere (Morikawa et al., 2004).
Results and Discussion
Figure 1 shows changes in the total leaf area of N. plumbaginifolia plants grown for 10 wk under natural light and irrigation with medium containing 10 mm KNO3 in air containing (+NO2 plants) or not containing (–NO2 plants) 150 ± 50 ppb 15NO2. During the first 5 wk, total leaf area did not greatly differ between +NO2 and –NO2 plants. To our surprise, however, total leaf area was larger in +NO2 plants than in –NO2 plants from around the 6th week onwards, and was 1.8 times larger in +NO2 plants than in –NO2 plants at harvest (10 wk).
Figure 2 depicts typical photographs of +NO2 plants (right) and –NO2 plants (left). Clearly, +NO2 plants exhibited a healthy green colour. The leaves and above-ground parts of +NO2 plants were markedly larger than those of –NO2 plants. The total number of leaves on an individual plant was slightly higher for +NO2 plants than for –NO2 plants; 19 ± 1 and 18 ± 1, respectively [mean ± standard deviation (SD) of six plants]. The difference in the size of the underground part of the plant (the root) between +NO2 and –NO2 plants was smaller (see Fig. 2).
Figure 3 summarizes shoot biomass, total leaf area, and contents per shoot of C, N, S, P, K, Ca, Mg, total free amino acids and crude proteins of +NO2 and –NO2 plants. Values for +NO2 plants (shaded columns in Fig. 3) are normalized relative to the respective value of 100 for –NO2 plants (open columns in Fig. 3). The shoot biomass, total leaf area, and contents per shoot of C, N, S, P, K, Ca, Mg, total free amino acids and crude proteins were 1.5–1.9 times greater in +NO2 plants than in –NO2 plants. Thus, +NO2 plants are proportionally similar to –NO2 plants (in their shoots) by a factor of 1.5–1.9 not only in size but also in the contents of major elements and metabolites. This means that, on a dry weight basis, the contents of C, N, S, P, K, Ca and Mg, free amino acids and proteins were almost the same for +NO2 and –NO2 plants. This is an indication that no biased accumulation of specific elements or metabolites occurred in +NO2 plants, suggesting that growth of +NO2 plants was ‘normal’. A histological investigation of the size and number of leaf cells is needed to clarify the mechanism by which +NO2 plants increased both leaf size and biomass (through cell enlargement or an increase in cell number, or both).
Because in the present study 15N-labelled NO2 and nonlabelled nitrate were fed to the plants, the 15N/14N ratio gives a measure of the content of NO2-N in total plant N. In mass spectrometry (Morikawa et al., 2004), it was found that NO2-N comprised < 3% of total N both in the shoots and in the roots of +NO2 plants; 3.0 ± 0.1% and 2.5 ± 0.1% (mean of three plants ± SD), respectively. This is a clear indication that the contribution of NO2-N to total N in +NO2 plants was negligible. It thus seems very likely that the primary role of NO2 is a multifunctional signal to stimulate plant growth, nutrient uptake and metabolism. We therefore propose that atmospheric NO2 at an ambient concentration may be a plant vitalization signal to double plant size and the contents of cell constituents.
Our present finding of the plant vitalization signal effect of exogenously supplied atmospheric NO2 shows that atmospheric NO2 is not a simple pollutant but instead acts as an airborne, ‘naturally occurring’ biological signal. To our knowledge, this is the first report of a gaseous material in the ambient atmosphere functioning as a signal in a living organism.
Plants do emit NOx (Wildt et al., 1997; Hari et al., 2003), and endogenously produced NOx reportedly functions as a multifunctional reactive nitrogen species (RNS) signal (Wendehenne et al., 2001; Neill et al., 2003 and references therein; Sakamoto et al., 2002, 2003, 2004) regulating multiple plant processes, including plant growth, differentiation, defence reactions and photomorphogenesis. However, whether exogenously supplied atmospheric NOx has a signal effect similar to that of endogenously produced NOx remained unclear. The present study shows for the first time that this is indeed the case, at least for NO2. Whether NO (another component of NOx) at an ambient concentration has the same biological effect as NO2 remains to be investigated.
As reported recently, about one-third of NO2-N (Morikawa et al., 2004) or nitrate-N (Morikawa et al., 2005) taken up into plants is metabolized in an alternative N pathway to form the UN. The UN-bearing compounds appear to include a novel heterocyclic Δ21,2,3 thiadiazoline derivative, and nitroso- and nitro-organic compounds (Miyawaki et al., 2004; Morikawa et al., 2005). It is possible that some of these novel N metabolites, if not all, are involved in a plant vitalization signal effect of atmospheric NO2.
This work was supported in part by a Grant-in-Aid for Scientific Research (no. 16208033) from the Japan Society for the Promotion of Science.