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Keywords:

  • nitrogen;
  • uptake;
  • foliar

Abstract

  1. Top of page
  2. Abstract
  3. 1. Introduction
  4. 2. Experimental
  5. 3. Results
  6. 4. Discussion
  7. Acknowledgments
  8. References

[1] Increasing nitrogen deposition to forests can impact the balance between the carbon and nitrogen cycles. This nitrogen source, if taken up and used by forests, can increase growth and carbon storage. While previous findings have suggested that nitrogen deposition is not an important source of nitrogen for a forest ecosystem, the possibility of canopy uptake was not considered. Foliar uptake and utilization of inorganic nitrogen, such as NO2 and NH3, has been shown to occur, but utilization of organic nitrogen has not been demonstrated directly. Here we show for the first time that atmospheric organic nitrates (RONO2), one form of organic nitrogen, can be taken up by foliage and incorporated into the leaf amino acids, and we discuss possible uptake mechanisms.

1. Introduction

  1. Top of page
  2. Abstract
  3. 1. Introduction
  4. 2. Experimental
  5. 3. Results
  6. 4. Discussion
  7. Acknowledgments
  8. References

[2] Increasing atmospheric carbon dioxide (CO2) concentration can increase photosynthesis and carbon storage in forests [Ollinger et al., 2002; Luo et al., 2004; Reich et al., 2006]. However, most forests are nitrogen limited, effectively constraining any CO2 fertilization effect [Townsend et al., 1996; Vitousek et al., 1997; Ollinger et al., 2002; Reich et al., 2006; Elser et al., 2007; LeBauer and Treseder, 2008]. Increased uptake of nitrogen from the soil, caused by increased CO2 uptake, depletes the amount of nitrogen available, depending on N pool sizes and residence times, unless a new source of N is added to the system [Luo et al., 2004]. Schimel and Bennett [2004, and references therein] have suggested that in low N environments, plants can compete with soil microbes for available nitrogen and can use organic N, not just inorganic N, as their N source. Deposition of anthropogenically derived nitrogen compounds, such as NO, NO2, HNO3, NH3, and aerosol nitrogen, along with organic nitrogen compounds such as PANs (peroxyacyl nitrates, RC(O)OONO2) and organic nitrates (RONO2) to the forest can increase nitrogen availability and thus potential ecosystem carbon storage [Gessler et al., 2002; Neff et al., 2002].

[3] There is some debate as to whether nitrogen deposition in any form is an important source of nitrogen to the forest. Nadelhoffer et al. [1999] suggested that atmospheric nitrogen deposition is not a significant source, but they did not include uptake by the canopy [Sievering, 1999]. In contrast, Sievering et al. [2000] concluded that about 90% of the deposited inorganic nitrogen undergoes uptake by the forest canopy, although that was not measured. Goodale et al. [2002], however, suggested that nitrogen deposited to vegetation might not be taken up and used. Nevertheless 15N labeling experiments have indicated that NO, NO2, HNO3, and NH3 can be taken up into the leaves and incorporated into plants [Vose and Swank, 1990; Hanson and Garten, 1992; Weber et al., 1995; Sparks et al., 2001; Gessler et al., 2002; Yoneyama et al., 2003]. Additionally, peroxyacetyl nitrate, comprising 12–30% of the reactive nitrogen (NOy) in the atmosphere (NOy = HNO3 + PANs + RO2NO2 + RONO2 + HONO + NO + NO2 + N2O5 + NO3-aerosol + NO2-aerosol + …) [Parrish et al., 1993; Murphy et al., 2006], has recently been shown to be taken up [Sparks et al., 2003], but has not yet been proven to be utilized by plants. Organic nitrates, which are chemically distinct from and more stable than PAN compounds, are produced from the atmospheric oxidation of volatile organic compounds (VOCs), such as isoprene (2-methyl-1,3-butadiene), the dominant biogenic VOC emission to the atmosphere [Guenther et al., 2006]. They comprise up to 20% of NOy and could be an important source of nitrogen for plants [Day et al., 2003; Murphy et al., 2006], but their uptake has not been studied. Here we show, by analysis of bulk leaf tissue and compound-specific stable 15N isotope analysis of leaf amino acids, that organic nitrates can be taken up and utilized by the foliage with direct incorporation into plant amino acids.

2. Experimental

  1. Top of page
  2. Abstract
  3. 1. Introduction
  4. 2. Experimental
  5. 3. Results
  6. 4. Discussion
  7. Acknowledgments
  8. References

[4] Leaves attached to one of four healthy 2 year old trembling aspen seedlings, Populus tremuloides (Greenwood Nursery, McMinnville, TN), were fumigated with a synthesized 15N-labeled organic nitrate, 1-nitroxy-3-methyl butane (MB15N, 99% purity, 99 atom% 15N), to determine whether organic nitrates could be taken up and utilized by the leaves. Aspen are fast growing deciduous trees native to most of the lower 48 US states (with the exception of the southeast), Alaska and Canada, and are substantial isoprene emitters [Guenther et al., 2006]. MB15N was chosen as a surrogate for the organic nitrates produced from the oxidation of atmospheric isoprene (2-methyl-1,3-butadiene). The trembling aspen trees were grown in Scotts Metro-Mix® Soil (366-P) in the Purdue University greenhouses (West Lafayette, IN) and kept at 24°C during the day and 18°C at night. Two-600 watt high pressure sodium (HPS) lamps (Sunlight Supply, Inc, Vancouver, WA) provided approximately 100 μmol m−2s−1 of photosynthetically active radiation (PAR) for 16 hours per day to supplement the natural sunlight in the greenhouse. The trees were watered using non-fertilized and fertilized water on alternating days. Nutrients were supplied from 1000 mg L−1 15-5-15 commercial fertilizer formulation (Miracle Gro© Excel© Cal-Mag; The Scotts Co., Marysville, OH).

[5] Two experiments were conducted to determine whether MB15N could be taken up and utilized. Attached trembling aspen leaves from each tree were fumigated with MB15N (350–700 parts per billion (ppb)) using a LI-COR 6400 flow-through leaf cuvette (LI-COR Biosciences, Lincoln, NE). The high MB15N concentration, which is much greater than ambient concentrations of organic nitrates, was chosen to approximate the total organic nitrogen leaf dose integrated over a growing season, assuming that the total concentration of atmospheric organic nitrates and PANs is between 0.4–1 ppb [Thornberry et al., 2001] and that the growing season is typically 150 days. The MB15N was added upstream of the leaf cuvette at a flow rate of 1 mL min−1 using a diffusion cell that was heated to 30°C (Figure 1). All of the sample lines after the addition of MB15N were maintained at 68°C to avoid surface losses. The relative humidity was maintained at 60% by passing the input air through a bubbler containing ultra pure water (Millipore, Bedford, MA) and then diluting that with zero air (Whitmore Clean Air Generator, Parker-Hannifin, Tewksbury, MA) using a mass flow controller (MFC).

image

Figure 1. Leaf cuvette experimental setup. The dark black lines are Teflon tubing that is heated to 68°C to avoid sample loss to the walls. The arrows represent the direction of the air flow through the setup. The cross by the bubbler represents the needle valve used to control the dilution flow for humidification. MFC stands for mass flow controller.

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[6] The intent for these experiments was to conduct replicate experiments under constant conditions. The CO2 concentration was 390 parts per million (ppm), PAR was 1000 μmol m−2s−1, the leaf cuvette temperature was 25°C, and the flow rate through the LI-COR was 300 mL min−1. The LI-COR 6400 measured photosynthesis, CO2 concentration, temperature, stomatal conductance, and relative humidity. The addition of the 1 mL min−1 of MB15N to the sample air flow after the LI-COR console did not affect the photosynthesis and conductance measurements. The MB15N concentration was measured before, during, and after the exposure using a HP 5890 Series II gas chromatograph (GC, isothermal at 150°C, RTX-1701 column, Restek) coupled with an electron capture detector (Valco, Houston, TX). The total amount of MB15N taken up by the leaf over the entire experiment was determined by subtracting the MB15N concentration when the leaf was in the cuvette from the concentration when the leaf was out of the cuvette and multiplying by the flow rate and total exposure time.

[7] The trembling aspen leaves used for the first experiment were washed with ultra pure water (Millipore, Bedford, MA), and dried in a 60°C oven to remove any leaf-surface adsorbed MB15N, then crushed and analyzed with a SerCon Ltd. elemental analyzer (CN-1) coupled to a PDZ-Europa 20/20 isotope ratio mass spectrometer (EA-IRMS, Cheshire, UK) at Purdue University's Stable Isotope Lab (West Lafayette, IN) to determine the total leaf δ15N value. For the second experiment, the amino acids were extracted from the trembling aspen leaves according to Rhodes et al. [1981], derivatized to their trifluoroacetyl isopropyl (TFA-IP) esters according to Silfer et al. [1991], and analyzed using a Shimadzu 17A GC (Columbia, MD) coupled to a PDZ-Europa 20/20 isotope ratio mass spectrometer (GC-IRMS, Cheshire, UK) at Purdue University's Stable Isotope Lab, according to procedures outlined by Macko et al. [1997] and Hofmann et al. [2003].

3. Results

  1. Top of page
  2. Abstract
  3. 1. Introduction
  4. 2. Experimental
  5. 3. Results
  6. 4. Discussion
  7. Acknowledgments
  8. References

[8] In the first experiment, one leaf was exposed to MB15N for 6.5 hours to determine total incorporation of 15N into the whole leaf, as δ15N (o/oo) (equation 1). One non-exposed leaf was used as the δ15N blank. Four replicate samples of the exposed leaf and three replicate samples of the blank leaf were analyzed using EA-IRMS. The δ15N values for the whole leaf experiment increased significantly to δ15N = 25.7 (±2, 1 s.d.) (all errors are ±1 s.d. unless stated otherwise)

  • equation image

relative to the blank (δ15N = −0.83 (± 0.38)), indicating that the 15N label was taken up into the leaf. However, this experiment does not address the question of foliar utilization of that nitrogen.

[9] To determine conclusively whether the 15N was utilized by the leaf, twelve trembling aspen leaves, 3 each from 4 separate trees, were exposed to MB15N for 8 hours and the δ15N values of glutamate (glu) and aspartate (asp) in the leaves were subsequently determined using GC-IRMS. Glu and asp were selected because known biochemical pathways indicate that glu would be the first amino acid formed from the labeled substrate and that asp would be formed by downstream biochemical processing in the leaf [Yoneyama et al., 2003]. The δ15N values of glu and asp of the exposed leaves were statistically different from the non-exposed leaves at the 95% confidence level (95% c.l.), indicating conclusively that the 15N label was taken up and incorporated into the leaf amino acids (Figure 2).

image

Figure 2. Glutamate and aspartate δ15N. The open bars are glutamate and the grey bars are aspartate. Each set of bars represents one leaf. The three leaves from each tree are grouped together and labelled to illustrate differences in δ15N between trees. The first set of bars is the average of all of the blank leaves for glutamate and aspartate. All samples were statistically significantly different from the blanks at 95% c.l. (student's t-test). The error bars for the samples are not visible; on average, the uncertainty is 0.5% (s.e.m.). The circles are the average leaf stomatal conductance (mmol m−2s−1). The error bars are the 95% c.l. (student's t-test).

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4. Discussion

  1. Top of page
  2. Abstract
  3. 1. Introduction
  4. 2. Experimental
  5. 3. Results
  6. 4. Discussion
  7. Acknowledgments
  8. References

[10] The potential impact of organic nitrate uptake can be determined using the average measured leaf MB15N uptake rate, 0.012 nmol 15N m−2 s−1 ppb−1 (±0.011). The average uptake rate was calculated using the difference in MB15N concentration when the leaf was in and out of the cuvette, integrated over the 8 hour exposure period, per area of leaf (m2) per exposure concentration (ppb). The large standard deviation in the observed uptake rate is not the measurement uncertainty, but the variability between experiments, which likely resulted from variations in leaf physiology and biochemistry, amino acid pool size, and exposure concentration, all of which could affect leaf uptake [Hanson and Garten, 1992; Peterson et al., 1999; Gessler et al., 2000; Sparks et al., 2001; Zak et al., 2004]. While the organic nitrate uptake rate is much lower than the reported foliar uptake rates for NO2 (0.03 to 0.31 nmol m−2s−1ppb−1) [Sparks et al., 2001] and PAN (0.15 nmol m−2s−1ppb−1) [Sparks et al., 2003], the trees were grown in N-rich soil, suggesting that the uptake rate might be higher than what was measured in this study compared to a real forested environment that is nitrogen-limited. Brukner et al. [1993] found that Norway spruce (Picea abies) trees grown in low nitrogen sand had a higher needle uptake of gaseous 15NH3 than trees grown in higher nitrogen sand, which suggests that foliar uptake of organic nitrates might be greater, but more experiments are needed to quantify the soil N-dependence of uptake.

[11] The uptake mechanism of MB15N is unknown, although there is information in our experiments about some of the physical aspects. Organic nitrates are not particularly chemically reactive [Roberts, 1990] and are only slightly soluble in water (Henry's Law constant is 0.74–1.6 M/atm at 295K for similar nitrates) [Roberts, 1990]. They hydrolyze in acidic or neutral solutions at a rate of 10−5 to 10−3 s−1 at room temperature [Robertson et al., 1981; Kames and Schurath, 1992]. There are 3 possible pathways for hydrolysis of organic nitrates [Baker and Easty, 1950; Connon, 1970]. The first is by nucleophilic attack on the carbon or nitrogen (reaction 2) to produce nitrate. The second is by abstraction of an α-hydrogen to produce nitrite (reaction 3), and the third is by abstraction of a β-hydrogen (dehydration reaction) (reaction 4), to also yield nitrate.The product NO3 or NO2 ions can then be reduced by nitrate/nitrite reductase to NH4+ and introduced into the biochemical cycle. If these pathways are responsible for the nitrogen utilization, it is likely that the rates are relatively small due to the small Henry's Law constants for solubilization in water.

  • equation image
  • equation image
  • equation image

[12] Although organic nitrates are rather lipophilic, it is reasonable to assume that, given the leaf washing and drying procedures for the first experiment, uptake occurred through the stomata rather than through the cuticle. Stomatal uptake of the gaseous MB15N is more likely as in the case of PAN uptake [Sparks et al., 2001]. However, the MB15N experimental conditions were such that the stomatal conductance for each experiment only varied by a factor of two (Figure 2). As a consequence, the data do not allow for evaluation of uptake rates as a function of stomatal conductance. The MB15N measured uptake accounted for 1.6–57% of the predicted uptake based on the modelled leaf stomatal conductance to MB15N, suggesting that there is an internal resistance to MB15N [Gessler et al., 2002; Teklemariam and Sparks, 2004], presumably, dissolution and hydrolysis. For comparison, the measured uptake for NH3 and NO2 by spruce needles was found to be greater than the predicted uptake, suggesting that there were additional sinks such as cuticular absorption and nitrifying bacteria on the surface and within the apoplast of the needles [Gessler et al., 2002; Papen et al., 2002]. Whether bacteria may have influenced the uptake of MB15N in these experiments is unclear.

[13] While the utilization mechanism for MB15N could occur via one of the hydrolytic mechanisms shown above, it is also possible that it occurred via an enzymatic or microbial process, possibly beginning with the breakdown of MB15N by endophytic fungi and bacteria. They are located within the leaf cells and intercellular space and are present in all temperate trees [e.g., Sieber, 2007]. The fungi and/or bacteria could use the carbon or convert it into secondary metabolites, such as phenols, steroids, and terpenoids that can benefit the tree, while releasing the nitrogen for use by the tree in a symbiotic relationship [Tan and Zou, 2001]. This has not been proven, but there is evidence that aerial fungi could behave similarly to mycorrhizal fungi, which provide the tree with much needed nutrients including nitrogen [Wilson, 2000; Schulz and Boyle, 2005, and references therein].

[14] These measurements add to the growing body of information that indicates that a wide range of N-containing atmospheric species of importance can be utilized by trees via stomatal or cuticular uptake and incorporation of the nitrogen into leaf amino acids. For some forests that are impacted by human inputs via atmospheric nitrogen pollution, this foliar uptake could be significant and requires further study. As forests become increasingly anthropogenically impacted, this N source may increase in importance and compensate for the increased atmospheric CO2 being emitted into the atmosphere by anthropogenic processes.

Acknowledgments

  1. Top of page
  2. Abstract
  3. 1. Introduction
  4. 2. Experimental
  5. 3. Results
  6. 4. Discussion
  7. Acknowledgments
  8. References

[15] We would like to thank Rob Eddy at Purdue University's greenhouses for his help in growing the trembling aspen trees, Sergey Oleynik at Purdue University's Stable Isotope lab for his help with the instrumentation, and the Biosphere Atmosphere Research and Training (BART) program at UMBS for the use of the LI-COR 6400 photosynthesis instrument. This work is based upon work supported by the National Science Foundation grant ATM0542701 and is paper number 0712 from the Purdue Climate Change Research Center.

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  2. Abstract
  3. 1. Introduction
  4. 2. Experimental
  5. 3. Results
  6. 4. Discussion
  7. Acknowledgments
  8. References
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