INTRODUCTION

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

Nitrate (NO₃) reduction in plants is an energy-intensive process, requiring eight high-energy electrons (e) for its reduction to ammonium (NH₄⁺), plus an additional two e for the Glu synthase activity needed to synthesize an amino acid, resulting in a total demand for 10 e.

In photosynthetic tissues, these electrons can be provided by the light reactions of photosynthesis and are therefore associated with the production of O₂ from water splitting and the subsequent use of the electrons for NO₃ reduction rather than for CO₂ fixation (Elrifi and Turpin, 1986; Bloom et al., 1989; Holmes et al., 1989). In non-photosynthetic tissues, the electrons come from carbohydrate metabolism and therefore are coupled to the production of CO₂, but the reducing power is not subsequently passed through the electron transport chain to consume O₂ (Abrol et al., 1983; Bloom et al., 1989, 1992; Huppe and Turpin, 1994).

Differences in the absolute values for rates of CO₂ (CER) and O₂ (OER) exchange have been associated with NO₃ reduction in Selenastrum minutum (Weger and Turpin, 1989), Dianthus caryophyllus (Avelange et al., 1991), Hordeum vulgare (Bloom et al., 1989, 1992; de La Torre et al., 1991), Pisum sativum (de La Torre et al., 1991) and Triticum aestivum (Bloom et al., 2002). However, these studies were limited to a qualitative correlation of gas exchange and NO₃ assimilation. They did not provide a quantitative analysis of the relationship between NO₃ assimilation and plant gas exchange that would allow the use of CER and OER measurements to provide a noninvasive measure of in situ NO₃ reduction in plant tissues. Such a quantitative analysis cannot be done from gas exchange ratios (e.g. Weger and Turpin, 1989; Bloom et al., 1992, 2002) or with information on percentage of changes of one gas relative to another (e.g. Bloom et al., 1989; Avelange et al., 1991). Rather, the measurements of CER and OER must be used to provide a measure of the rate of electron flow to sinks that do not involve the uptake of CO₂ or O₂.

The term diverted reductant utilization rate (DRUR = 4*[CER+OER]; where gas production is positive, uptake is negative; units of moles of e per hour) has been defined as a measure of the rate of electron flow to biosynthetic processes that result in the synthesis of biomass that is more reduced (positive DRUR) or more oxidized (negative DRUR) per unit C than the initial biomass in respiring tissues or than carbohydrate in photosynthesizing tissues (Willms et al., 1999; Cen et al., 2001). The "4" in the above equation refers to the four e associated with one CO₂ that is produced by carbohydrate catabolism, one O₂ that is produced in the light reactions of photosynthesis, one O₂ that is taken up in the respiratory electron transport chain, or one CO₂ that is fixed in the dark reactions of photosynthesis.

The use of DRUR as a measure of NO₃ reduction in plant tissues must also take into account the effects that NO₃ assimilation will have on C metabolism, especially the production of the C skeletons used in synthesizing amino acids. For example, the conversion of Glc to the oxoglutarate needed for the synthesis of Glu would be associated with the production of two CO₂ molecules from the Kreb's cycle, the generation of eight e that are presumably coupled to two O₂ uptake, and the fixation of one CO₂ resulted through phosphoenolpyruvate carboxylase. The net gas exchange (CER + OER) of this pathway would be equivalent to 1, yielding a DRUR equivalent of 4 e per NO₃ assimilated. When this is added to the reductant cost of NO₃ assimilation and GOGAT activity (10 e per NO₃ assimilated), the apparent reductant cost would be 6 e per NO₃ assimilated.

The C skeletons of other amino acids may be more or less oxidized than Glu, resulting in predicted values for the apparent reductant cost of NO₃ assimilation that are less than or greater than, respectively, that for Glu. For example, Asp synthesis from Glc and NO₃ would have an apparent reductant cost of 4 e per NO₃ assimilated, whereas Asn, Gln, and Pro synthesis would have apparent reductant costs of 6, 7, and 10 e per NO₃ assimilated, respectively (calculations not shown).

The first whole-plant measurement of DRUR was reported in white lupin (Lupinus albus; Cen et al., 2001), and an attempt was made to relate these to the measured growth and likely composition of the new biomass that was produced during the period of measurement. Assuming a standard composition for the newly formed biomass (Cen et al., 2001) and 4 to 10 electrons consumed per NO₃ assimilated, 61% to 80% of the DRUR should be attributed to NO₃ assimilation. The remaining 20% to 39% would be associated with the synthesis of lipids or other organic molecules that are more reduced per unit C than carbohydrates (calculations not shown).

If it is possible to provide a quantitative, experimentally supported link between measured DRUR and NO₃ assimilation in plant tissues, a valuable noninvasive tool could be developed to address long-standing questions regarding the site, timing, and regulation of NO₃ assimilation in plant tissues. For example, in soybean (Glycine max), NO₃ reduction has been reported to be either predominantly shoot-based (McClure and Israel, 1979; Rufty et al., 1982; Andrews et al., 1984; Rufty et al., 1984; Andrews, 1986) or predominantly root-based (Radin, 1978; Crafts-Brandner and Harper, 1982; Vessey and Layzell, 1987). Although these discrepancies may be attributed to differences in cultivar or experimental conditions, there are also major differences among studies in the assay techniques that are used to measure the site of NO₃ assimilation.

The present study explores the use of CER and OER as a noninvasive measure of NO₃ reduction in roots and shoots of soybean (cv Maple glen) by comparing the difference in plant DRUR between plus and minus NO₃ with ¹⁵NO₃ assimilation. It was hypothesized that NO₃ assimilation should account for about 61% to 80% of the whole-plant DRUR in vegetatively grown soybean plants and have a reductant cost of between 4 to 10 mol e mol¹ NO₃ assimilated.

	RESULTS

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

Continuous CO₂ and O₂ Measurements

A gas exchange system was designed and built that permitted the simultaneous measurement of CER and OER in the roots and in the shoot tissues (leaf or stem + petiole) of soybean plants. Figure 1 shows example data for the CO₂ and O₂ concentration in a reference (inlet) gas stream (channel R), a calibration gas stream (channel C), and the eight effluent gas streams from pots containing soybean roots provided with NO₃ (channels 1-8). A hand-held leaf cuvette was incorporated into a parallel gas exchange system, making it possible for rapid (4-10 min leaf¹) analysis of CO₂ and O₂ exchange under steady-state conditions. This approach made it feasible to study simultaneous CO₂ and O₂ exchange in a single leaf with time or among a population of leaves that vary in physiological or environmental conditions. To our knowledge, this is the first system to incorporate simultaneous CO₂ and O₂ measurements using a small, handheld leaf chamber.

View larger version (33K): [in this window] [in a new window]   Figure 1.   A complete cycle of gas exchange measurements obtained from intact root systems of NO3-grown soybean plants. Channels 1 to 8 are measurements of the effluent gas streams from the pots of eight replicate plants, whereas channel R is the reference or inflow gas stream, and channel C is a calibration gas stream. The voltage signals were converted to pascals of CO2 or O2 before plotting. The CO2 values were absolute measurements when the reference gas stream was CO2-free air, and the O2 values represent the differences between the reference (about 21,000 Pa O2) and the sample or calibration gas streams.

To compensate for drift in the sensitivity of the sensors over the measurement period, calibration gas streams having a known ratio of CO₂ enrichment (or depletion) and O₂ depletion (or enrichment) were prepared and assayed every 40 min (Willms et al., 1997, 1999; Cen et al., 2001). Over a 4-d period, less than 2% variation was observed in the relative response of the infrared CO₂ gas analyzer (IRGA) and differential O₂ analyzer (DOX) instruments when presented with the calibration gas stream (data not shown). Therefore, the calibration system provided confidence in the quality and reliability of the data sets that were obtained.

The Effect of N Supply on Reductive Metabolism of Roots

Root gas exchange was measured on a whole-plant basis, but presented as percent of initial CER to remove plant-to-plant variability and make it easier to identify diurnal patterns (Fig. 2A). CER values were positive at all times, but peaked at about 6 h into the light period and again at the end of the light period (Fig. 2A, solid line). Over the 8-h dark period, CER declined. Values for OER were negative and mirrored the trends observed in CER; however, the absolute values for OER were lower than those for CER in all root systems at all times. When simultaneous measurements of CER and OER were converted to dry-weight specific values (from the root biomass), they were used to calculate DRUR. A peak in DRUR was observed in the middle of the light period, and a trough was observed in the middle of the dark period (Fig. 2C, solid line). This pattern was repeated over the 3-d measurement period, and on d 25, the average DRUR was about 25 nmol e g¹ dry weight s¹, with 26 and 22 nmol e g¹ dry weight s¹ in the light and dark, respectively.

View larger version (40K): [in this window] [in a new window]   Figure 2.   The CER and OER (A and B) and the DRUR (C and D) of roots of soybean plants that received nutrient solution containing 5 mM NO3 (N+, solid lines) and 0 mM NO3 (N, dashed lines). A and C show data for plants that were grown with N+ nutrients before a N treatment (arrows). B and D were pretreated with N for 2 d before a N+ treatment (arrows). The shaded regions show the timing of the 8-h dark periods. The vertical bars were ±SEs, n = 4. Values in A and B were expressed as percentage of initial CER, whereas those in C and D were normalized to gram of root dry weight.

On d 24, after 1 d of measurements, some of the plants were flushed with a NO₃-free nutrient solution (N treatment; Fig. 2, A and C, arrow). This significantly lowered the root CER and made the OER less negative, but the overall diurnal patterns remained unchanged (Fig. 2, A and C, dashed line). However, the absence of NO₃ in the nutrient solution altered CER more than OER, leading to a sharp decline in the DRUR in the roots of the N plants. By d 25, the DRUR in the N plants was about 25% of that in the 5 mM NO₃-feed (N+ treatment) plants, and the magnitude of variations between the light and dark period was much lower (Fig. 2C, dashed line).

When the N plants were subsequently watered with a N+ nutrient solution on d 26 (N± treatment), the CER was increased, and OER became more negative relative to the N plants (Fig. 2B, solid line). However, the changes in CER were more pronounced than OER, leading to a significant increase in DRUR by d 27 (Fig. 2D, solid line).

The Effect of N Supply on Reductive Metabolism of Stem + Petioles

Simultaneous CER and OER measurements were made in the dark on excised stem + petioles from plants harvested during the latter part of the dark period and early part of the light period. The values were normalized to specific activities using the dry weights obtained at the end of each 24-h measurement period. No significant differences were observed in gas exchange rates of tissues given the same N treatment (data not shown), so values were combined for presentation in Figure 3. In the N+-treated plants, the CER values were greater than the absolute values of OER (Fig. 3A), leading to a DRUR of about 3.9 nmol e g¹ dry weight s¹ (Fig. 3C), a value that was about 5-fold less than that obtained for an equivalent weight of roots from the same plants. Significantly lower DRUR (about 1.5 nmol e g¹ dry weight s¹) were found in plants provided with the N solution (Fig. 3C). When the N plants were treated with a N+ nutrient solution again, the DRUR increased within 24 h (Fig. 3D).

View larger version (36K): [in this window] [in a new window]   Figure 3.   The CER and OER (A and B) and the DRUR (C and D) of stem + petioles of soybean plants that received nutrient solution containing 5 mM NO3 (N+, white boxes, solid lines) and 0 mM NO3 (N, shaded boxes, dashed lines). A and C, Data for plants grown with N+ nutrients before a N treatment (arrows). B and D, Pretreated with N for 2 d before a N+ treatment (arrows). The shaded regions show the timing of the 8-h dark periods. The vertical bars were ±SEs, n = 8.

The Effect of N Supply on Reductive Metabolism of Leaves

In the leaves, gas exchange measurements were made and presented on a leaf area basis, but converted to a gram dry weight basis using the measured relationship before calculation of DRUR. In N+ grown plants, the rates of gas production and consumption in the light were about 6-fold higher compared with the dark period (Fig. 4A). In the light, the leaves had a positive OER that was 0.1 to 0.3 µmol m² s¹ greater than the absolute value of a negative CER, but in the dark, a positive CER was obtained that was 0.08 to 0.12 µmol m² s¹ greater than the negative OER (Fig. 4, A and B). Therefore, on a gram dry weight basis, the DRUR of leaves averaged 18 nmol e g¹ dry weight s¹ for a 24-h period with values ranging from 22 nmol e g¹ dry weight s¹ in the light period to 10 nmol e g¹ dry weight s¹ in the dark. The leaves of N+ soybean had dry weight-specific DRUR values that were similar to that of roots but about 4-fold greater than that of stem + petioles.

View larger version (40K): [in this window] [in a new window]   Figure 4.   The CER and OER exchange rates (A and B) and the DRUR (C and D) of leaves of soybean plants that received nutrient solution containing 5 mM NO3 (N+, white boxes, open or hatched) and 0 mM NO3 (N, shaded boxes, open or hatched). A and C, Data for plants that were grown with N+ nutrients before a N treatment (arrows). B and D, Pretreated with N for 2 d before a N+ treatment (arrows). The shaded regions show the timing of the 8-h dark periods. Note that the photosynthetic measurements during the light period (left scale) were plotted on a different scale than the respiration measurements during the dark period (right scale). The vertical bars were ±SEs, n = 6.

When the plants were given an N nutrient solution, there was little effect compared with the N+ treatment on the gas exchange in the subsequent dark period. However, after 24 h without N (on d 25), the leaves showed reduced CER and less negative OER, resulting in DRUR values that were significantly lower than that in the plants provided with N+ nutrients (Fig. 4C). When the N-treated plants were provided with N+ nutrients, a significant increase was observed in OER and DRUR in the light period within 24 h (Fig. 4, B and D).

NO₃-Associated DRUR and ¹⁵N Accumulation in Soybean

To calculate the DRUR associated with NO₃ assimilation on a whole-plant basis, the dry weight-specific values for the N+ and N treatment over d 25 (Figs. 2C, 3C, and 4C) were multiplied by the grams dry weight on d 26 (Table I) to return the values to whole-plant rates. The difference in these DRUR values between the N+ and N treatment in roots (Fig. 2), stem + petioles (Fig. 3), and leaves (Fig. 4) were calculated as a measure of the DRUR associated with N reduction (DRUR_Nred, Eq. 1; see "Materials and Methods") for each plant organ for d 25. 

View this table: [in this window] [in a new window]   Table I.   The dry weights of soybean plants grown with (N+) and without (N) NO3 (5 mM) in the nutrient solution after 24 d of growth on N + nutrients Values are mean ± SE (n = 10).

The results (Fig. 5A) showed that during the 16-h light period, the accumulated DRUR_Nred was 4.56 mmol e plant¹, with 53% of this occurring in leaves, 39% in roots, and 8% in stem + petiole tissues. Over the subsequent 8-h dark period, an additional 1.02 mmol e plant¹ of DRUR_Nred was achieved for a total of 5.58 mmol e plant¹ over an entire day. However, during the dark period, most of DRUR_Nred was localized in root tissue (69%), although some occurred in stem + petiole tissues (18%) and in leaves (13%).

View larger version (18K): [in this window] [in a new window]   Figure 5.   A cumulative plot of the NO3-linked diverted reductant utilization (A) and the newly reduced N (B) that occurred in roots, stem + petioles (S+P) and leaves over the light and dark periods of d 25. Values for A were calculated as the difference between plants receiving 5 mM NO3 (DRURN+) and 0 mM NO3 (DRURN), whereas the accumulation of reduced N was calculated from assays of tissue 15N following the supply of plants with 50% abundance of 15NO3. Data in A and B were normalized to the dry weight of 25-d-old plants in Table I.

To obtain an independent measure of the actual rate of N uptake and assimilation, ¹⁵N accumulation was measured in plant tissues of a population of plants grown under identical conditions but provided with ¹⁵NO₃ (50% abundance of ¹⁵N). Assuming no changes in the relative pool sizes of NO₃ in plant organs (8.7%, 6.9%, and 3.9% of total N in roots, stem + petioles, and leaves, respectively; Table II), the measurements of ¹⁵N accumulation were used to calculate the N that was taken up, reduced, and incorporated into plant tissues (reduced N deposition) during the light and dark periods of 25-d-old plants. The dry weight-specific values for reduced N deposition in the plant parts of the ¹⁵NO₃-treated plants were multiplied by the dry weight values from corresponding parts of the 25-d-old plants that were used for gas exchange (Table I) to obtain a whole-plant value for reduced N deposition. Because the dry weight of the 25- to 26-d-old plants used in the ¹⁵N experiments (7.08 ± 0.25, Table II) was similar (P > 0.1, Student's t test) to that of plants used in the gas exchange experiments (6.90 ± 0.18, combined values of 25- and 26-d-old N+ plants; Table I), this correction was minor. These reduced N deposition values were then compared with those for DRUR for the 25-d-old plants having the same dry weight.

View this table: [in this window] [in a new window]   Table II.   The total nitrogen, NO3 content, and 15N accumulation in 25- to 26-d-old soybean plants provided with 5 mM 15NO3 (50/50 K15NO3 / KNO3) twice a day (2 h after light on and 1 h before light off) and harvested at time intervals from 0 to 24 h after the first exposure to 15N Values are means (if n = 2) or means ± SE of n replicate samples.

When the values for N reduction and incorporation were presented in a cumulative plot from the start of the ¹⁵N treatment, the results showed that a higher proportion (76%) of the whole-plant N assimilation occurred in the 16-h light period than in the subsequent 8-h dark period (24%; Fig. 5B). Of the N reduced during the light period, leaves, stem + petioles, and roots accounted for 61%, 17%, and 22%, respectively (Fig. 5B). Over the dark period, the reduced N in leaves, stem + petioles, and roots accounted for 32%, 20%, and 48%, respectively (Fig. 5B).

Integrated over the entire 25 d, 0.61 mmol NO₃ was taken up and 0.58 mmol was reduced by the whole plant, with the leaves, stem + petioles, and roots accounting for 54%, 18%, and 28%, respectively (Fig. 5B).

The whole-plant reduced N accumulation reflected NO₃ reduction at the plant level regardless of the site (shoot versus root) of NO₃ reduction. Because the DRUR_Nred represented the flow of reductant associated with N assimilation for 25-d-old plants, the measured reductant cost of NO₃ assimilation (_Nred) was calculated using Equation 4 (see "Materials and Methods"), to give a value of 9.6 mol e mol¹ N.

Shoot versus Root NO₃ Assimilation

The DRUR_Nred values for shoot (DRUR_Nred^Sht) and root (DRUR_Nred^Rt) were used in Equation 5a (see "Materials and Methods") to calculate the proportion of whole-plant NO₃ reduction (N_Red^Plant) that occurs in the shoot (N_Red^Sht; Fig. 6A). In 25-d-old soybean plants, 61%, 31%, and 55% of the whole-plant NO₃ assimilation was calculated to be in the shoots during the light, dark, and integrated over the day, respectively (Fig. 6A).

View larger version (15K): [in this window] [in a new window]   Figure 6.   NRedSht to NRedPlant calculated from measured values for the DRURNredRt and DRURNredSht (A) or from measurements of DRURNredRt and the whole-plant increment in reduced N by 15N (B). The experimental data for light (white circles), dark (black circles), and daily (gray circles) are shown for 25-d-old soybean plants.

When similar values were calculated using Equation 5b (see "Materials and Methods") with data on whole-plant N assimilation (from ¹⁵N) and values of DRUR_Nred^Rt, it was necessary to select a value for _Nred, the reductant cost of NO₃ assimilation. Using the measured value for _Nred of 9.6 mol e mol¹ N, the proportion of whole-plant NO₃ assimilation estimated to be in the shoot was 58%, 39%, and 55% in the light, dark, and integrated over the day, respectively (Fig. 6B).

	DISCUSSION

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

NO₃-Linked DRUR and the Reductant Cost of NO₃ Assimilation in Soybean

In the present study, N plants had daily whole-plant DRUR values that were 36% of N+ plants, but this proportion changed in light (31%) and dark (52%). Therefore, integrated over a 24-h period, the DRUR linked to NO₃ assimilation was 64% of whole-plant DRUR. This value was in the range of values predicted for a "typical" plant (Cen et al., 2001) assuming a reductant cost of NO₃ assimilation of 4 to 10 e per NO₃ assimilation.

Of course, the mineral and organic matter composition of the plants used in this study may have varied significantly from the typical plant described by Cen et al. (2001). Nevertheless, the experimental results obtained here and the theoretical calculations are consistent with N assimilation having the predominant role in determining the imbalance in CER and OER of plant tissues and the values of DRUR.

The measured reductant cost for NO₃ assimilation (_Nred = 9.6 mol e mol¹ N) was within the predicted range of 4 to 10 e per NO₃ assimilated, albeit on the higher end of the range. Despite the fit within the predicted range, too much confidence should not be placed in the 9.6 value because there may be a number of potential sources of error in the current study. For example, the plants used for gas exchange measurements were from a different population of plants than those used for the ¹⁵N studies. Both measurements should ideally be made simultaneously on the same plants. Also, the leaf gas exchange rates were measured with a small, hand-held cuvette, and there may have been a tendency to favor some of the larger leaves that were exposed to higher light levels, potentially overestimating actual whole-shoot gas exchange rates and the DRUR_Nred^Sht. On the other hand, there was good confidence in the actual measurements of DRUR of plant tissues because the simultaneous measurements of CER and OER exchange removed most of the common-mode noise, thereby allowing for measurement of DRUR differences between treatments in studies where there were no significant differences in actual measurements of CER or OER.

Using DRUR Measurements to Study Plant NO₃ Assimilation

Two strategies are presented for the use of plant CO₂ and O₂ gas exchanges in providing a quantitative measure of the spatial and temporal variations of plant NO₃ assimilation.

Method 1. Direct Measurement of DRUR in Shoots and Roots

Method 2. Coupled Measurements of Root DRUR_Nred and ¹⁵N Assimilation

NO₃ Assimilation in Soybean Plants

When the two methods were used to calculate the N_Red^Sht to N_Red^Plant ratios in the present study (Fig. 6), the results showed that neither roots nor shoots were the dominant site of NO₃ reduction in soybean. Rather, both organs shared the responsibility, with roots being proportionally more important during the dark (53%-69% of whole plant), whereas shoots were more important during the light (58%-61% of whole plant).

These light-dark differences of the sites of NO₃ assimilation on a single plant may explain the controversy in previous studies over whether the site of NO₃ reduction is in roots (Radin, 1978; Crafts-Brandner and Harper, 1982; Vessey and Layzell, 1987) or in shoots (Hatam and Hume, 1976; McClure and Israel, 1979; Rufty et al., 1982; Andrews et al., 1984; Andrews, 1986). Using the methods developed here, it would be interesting to explore how nutrient supply (NO₃ availability, atmospheric CO₂, and nodulation), environmental conditions (light, temperature), or plant ontogeny (vegetative versus reproductive) affects the relative balance between root and shoot NO₃ reduction in soybean. Once NO₃ is reduced in plant tissues, it can be translocated to other plant organs in either the xylem or phloem (Rufty et al., 1982; Vessey and Layzell, 1987). This explains why the organ distribution of NO₃ reduction measured as DRUR_Nred (Fig. 6) differs significantly from the distribution of reduced N deposition measured by ¹⁵NO₃ analysis (Fig. 5B).

However, by comparing these two data sets, it is possible to gain some insights into the relationships between NO₃ assimilation and transport in soybean. For example, over d 25, root NO₃ assimilation accounted for 45% to 52% of the whole-plant NO₃ assimilation, although roots accounted for only 28% of the whole-plant N deposition. The remainder of 17% to 24% of N reduced in root was transported to the shoot, presumably in the xylem. The xylem would also carry to the shoot 48% to 55% of the NO₃ taken up by the plant, of which 17% was reduced in the stem + petioles, and 83% in the leaves (Fig. 5A).

A more comprehensive study, perhaps combined with an analysis of xylem and phloem composition (Layzell and LaRue, 1982) in addition to DRUR_Nred and ¹⁵NO₃ measurements may permit a full assessment of the uptake, transport and assimilation of NO₃ and reduced N in plant tissues.

Coupling Plant Gas Exchange, NO₃ Assimilation and C Metabolism

The gas exchange method developed here for measurement of NO₃ assimilation in plants requires a better understanding of the relationship between the DRUR of plant tissues and the NO₃ assimilation rate in that tissue. Further development of this method would benefit from a detailed biochemical model linking tissue CO₂ and O₂ exchanges to the reductive and oxidative biosynthetic pathways that are occurring within those tissues. Given that NO₃ reduction to NH₄⁺ is the dominant pathway in plants that leads to imbalances in CO₂ and O₂ exchanges (Cen et al., 2001), it would also be necessary to explore how values for _Nred would be affected by changes that may occur in the pathways for the production or use of non-nitrogenous C compounds that may be coupled to NO₃ reduction. For example, shoot NO₃ reduction has been associated with the synthesis and phloem export of malate to maintain a pH and charge balance resulting from the release of OH during NO₃ reduction (Raven and Smith, 1976).

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

Plant Material and N Treatments

Seeds of soybean (Glycine max L. Merr. cv Maple glen) were sterilized (0.3% [w/v] sodium hypochlorite for 3 min) and rinsed more than four times with distilled water for at least 5 min. They were then planted in silica sand in 0.72-L pots that could be sealed for gas exchange measurements (Hunt et al., 1989). The plants were watered twice daily (2 h after the start, and 1 h before the end of the photoperiod) with a nutrient solution (Walsh et al., 1987) containing either 1 mM (0-10 d) or 5 mM KNO₃ (11 d+). They were maintained in a growth chamber (model PGV 36, Conviron, Winnipeg, MB, Canada) having a 16-h photoperiod and a temperature of 21°C. Photosynthetically active radiation was 500 µmol quanta m² s¹ at the plant level, and relative humidity was 75%. A population of 192 plants on four carts (48 plants each) was rotated 180^o daily and switched position with other carts twice a week to minimize location effects in the growth chamber.

After 24 d of growth, uniform plants (102) were randomly selected to receive one of the three treatments (34 plants per treatment). Some plants were maintained on 5 mM NO₃, others were transferred to a N-free nutrient solution on d 24, and the remainder received N-free nutrient solution on d 24 and 25, but were given 5 mM NO₃ on d 26 and 27.

Another population of plants was grown under identical conditions as the N+ treatment, but provided with 5 mM K¹⁵NO₃ (50% ¹⁵N enrichment, USDE, Monsanto Research Corporation, Mound Facility, Miamisburg, OH) at the usual watering times on d 25.

The Gas Exchange System

An open gas exchange system was built to provide measurements of CER and OER simultaneously in plant roots and in either leaves or stem + petioles (Fig. 7).

View larger version (47K): [in this window] [in a new window]   Figure 7.   Schematic diagram of the gas exchange system that was built for simultaneous multichannel measurement of CO2 and O2 exchange from eight root systems and a leaf or stem + petiole cuvette. The cm and p.s.i. values show the regulated pressure values throughout the system. See text for description. Air, Compressed air; R, pressure regulator; MV, mixing volume; A through K and S, flow meters for gas mixing; C1 through 3, flow meter for calibration gas streams; R1 through 5, flow meter for reference gas streams; Drying, magnesium perchlorate column; Soda lime, CO2 scrubbing column; V1 through 9, needle valves; V10 and 11, four way switching valve; DP, differential pressure sensor; and O2, O2 sensor.

The gas exchange system for roots used compressed air drawn from outside the building. The air was humidified and scrubbed of CO₂ using two soda lime columns (0.13 m³ each) before being provided to a bank of flow meters (Fig. 7, A, D, F, and G). The gas from flow meter F (Fig. 7) was distributed to the root system of eight pots (Fig. 7, 1-8) at a flow rate of about 300 mL min¹ for each pot and to three reference gas streams (Fig. 7, R 1-3). To provide a calibration gas stream (Fig. 7, C1) that was enriched in CO₂ and depleted in O₂ by a precisely defined ratio (Cen et al., 2001), the CO₂ free-air stream (Fig. 7, G) was mixed with a gas stream of 21% (v/v) CO₂ in N₂ (Fig. 7, V3).

The eight root samples, one reference, and the calibration gas streams were connected to a computer-controlled, multichannel gas sampling system (Layzell et al., 1989). This system selected one gas stream at a time (4 min per gas stream) to be delivered to a drying column (magnesium perchrorate, 15 mL), DOX (Fig. 7, DOX1; S-3A/DOX, AEI Technologies, Pittsburgh), and IRGA (Fig. 7, IRGA1; model S151, Qubit Systems Inc. Kingston, ON, Canada). A reference gas stream (Fig. 7, R2) was also dried and provided to the reference cell of the DOX.

A separate component of the gas exchange system was designed for single, short-term measurements of CER and OER in leaf or stem + petioles tissues. In this system, CO₂-free air (Fig. 7, D) was mixed with 21% (v/v) CO₂ in N₂ (Fig. 7, E) to provide a gas stream with a CO₂ concentration (typically 410-450 ppm) required to maintain CO₂ around a photosynthesizing leaf at about 360 ppm.

This CO₂-enriched gas stream was provided to a bank of flow meters (Fig. 7, B, C, S, and R4-5) at a background pressure of a 60-cm water column. Gas from flow meter C (Fig. 7) was mixed with 21% (v/v) CO₂ in N₂ (Fig. 7, V4) to provide a calibration gas stream (Fig. 7, C2) that was enriched in CO₂ and depleted in O₂ (respiration calibration). On the other hand, gas from flow meter B (Fig. 7) was mixed with CO₂-free air (Fig. 7, A) to provide calibration gas stream (Fig. 7, C3) that was enriched in O₂ and depleted in CO₂ (photosynthesis calibration).

A sample gas stream (S, 50-180 mL min¹) was provided to a Parkinson leaf cuvette (leaf area of 6.5 cm²; model PLC-B, Analytical Development, Hoddesdon, UK), and the pressure in the effluent stream was monitored by a manometer before being provided to two 4-way values that were used to select the stream sent for analysis (Fig. 7, V10-11). A bypass loop was used to maintain the humidity in the chamber (RH = 75%-80%) during CER and OER measurements. Other gas streams provided to the values included the two calibration streams (Fig. 7, C2 or C3) and a reference gas stream (Fig. 7, R4). The selected gas was provided to an IRGA (IRGA2; LI-6262, LI-COR Inc. Lincoln, NE) before being subsampled by a pump, and sent to a drying column (magnesium perchlorate, 15 mL) and DOX (Fig. 7, DOX2; S-3A/DOX, AEI Technologies). A CO₂-free and dried reference gas stream (Fig. 7, R3) was provided to the reference cell of IRGA2. A dried reference gas stream (Fig. 7, R5) with required composition of CO₂ was flow rate controlled by flow meter K (Fig. 7) and provided to the reference cell of DOX2.

To measure the CER and OER of stem + petioles, the Parkinson leaf chamber was replaced by a 60-mL syringe (BD Biosciences, Franklin Lakes, NJ) into which excised stem + petioles were placed. After 10 to 15 min, stable gas exchange rates were obtained at a flow rate about 200 mL min¹. Because light (150-300 µmol quanta m² s¹) had little effect on the measured rates of CER and OER exchange (data not shown), the measurements were assumed to reflect both daytime and nighttime activities.

Calculating CER and OER from CO₂ and O₂ Measurements

The voltage signals from the two DOX instruments included output signals for differential and absolute O₂ concentration, differential and absolute pressure, and temperature of the differential O₂ sensor block. These signals, and signals from the two IRGAs (Fig. 7, absolute CO₂ from IRGA1 and absolute CO₂ and absolute H₂O from IRGA2) were collected every 10 s using Workbench software (v4.01, Strawberry Tree Software, Sunnyvale, CA) running on a Macintosh computer (Apple Computer, Cupertino, CA). A complete cycle of root gas exchange measurements is shown in Figure 1. The dwell time on each channel was 4-min, and data for the last 100 s of each channel measurement were averaged for subsequent calculations.

The CER and OER of plant leaves were measured on attached leaves when steady-state gas exchange was achieved. A sample flow rate of about 180 and 60 mL min¹ in the light and dark, respectively, resulted in an O₂ differential between the inlet and outlet gas streams of about 6 Pa for photosynthesis and 3 Pa for respiration. Corrections for the effects of water vapor on CER of the leaves and stem + petioles were carried out according to von Caemmerer and Farquhar (1981) because the CO₂ was measured before dehumidification.

The differences between the inlet and effluent gas streams were combined with information on the gas flow rate through the chambers to calculate the CER and OER where positive and negative values were rates of gas production and uptake, respectively. These calculations considered the dilution effects on pO₂ associated with an imbalance in CO₂ and O₂ exchange (Willms et al., 1999). The CER and OER values were used to calculated the DRUR for each tissue as described previously (Cen et al., 2001).

Values for root measurements of gas exchange and DRUR were obtained every 40 min during the day and night, whereas measurement of leaf gas exchange were averaged to give a single daytime and a single nighttime value (four plants per treatment and three to six fully expanded leaves per plant) were measured during each daytime or nighttime period). In stem + petioles, measurements of four plants per treatment were averaged to give one value for each 24-h period.

To obtain a measure of the NO₃-linked DRUR (DRUR_Nred^X, moles of e per hour) for each plant part (X), the difference was calculated between the DRUR in the N+ treatment and the DRUR in the N treatment in 25-d-old plants. Therefore, for each light and dark period:

(1)

where X was the roots, stem + petioles or leaves, respectively.

Measuring the Uptake, Reduction, and Cost of NO₃ Assimilation

Plants for growth analysis were randomly selected from the N+, N, and N± treatments. After separating the leaves, stem + petioles, and roots, the whole-plant leaf area was estimated according to Cen and Bornman (1990), the plant tissues were dried in an oven (70°C) to constant weight (120 h), and then dry weights of the tissues were determined. In addition, plants randomly selected for gas exchange measurements were harvested at the end of the study period, and their biomass were determined. Finally, in the ¹⁵N treatment, six plants were harvested at 0, 2.5, 12, 16, and 24 h after exposure to K¹⁵NO₃.

The plant material from the ¹⁵N treatment was ground to a 1-mm mesh size, digested at 420°C with concentrated H₂SO₄ plus Kjeltabs catalyst (3.5 g K₂SO₄ per 3.5 mg Se) using a Kjelter System 1002 Distilling Unit (Tecator, Hoganas, Sweden) to determine total N (Zhou et al., 1998a). The atomic percent ¹⁵N of the plant sample was determined by emission spectrometry using a ¹⁵N analyzer (N-150, JASCO, Ltd., Tokyo) as described by Zhou et al. (1998b). The NO₃ contents in various plant tissues were determined by colorimetry as described previously (Cataldo et al., 1975).

The increase in the percent abundance of ¹⁵N between ¹⁵N applied sample plants (A_S) and control plants (A_O) for each plant parts was used to calculate the rate of NO₃ that was taken up and stored during the period (N_Uptake, moles per hour):

(2)

where X denoted roots, stem + petioles or leaves; N_Total was the total N content in grams per gram dry weight; DW was the dry weight in grams; A_S was the atomic percent of ¹⁵N in sample plants given ¹⁵NO₃ as the N source; A_O was the atomic percent of ¹⁵N of control plant; MW_15N was the M_r of ¹⁵N in grams per mole; MW_N was the M_r of N in grams per mole; C was the atomic percent of ¹⁵N source applied to plant (50%); H was the time between first application of ¹⁵NO₃ and the harvest in hours.

  To calculate the N that was reduced and stored in each plant part during this period (N_Red, mol h¹), the NO₃ concentration ([NO₃], percentage of total N) was measured in each plant part, and an assumption was made that the NO₃ pool size (ratio of NO₃ to total N) did not change over the course of the experiment. Therefore,

(3)

where X denotes the roots, stem + petioles, or leaves.

  The sum of N_Red^X for all plant parts (X) over d 25 provided a measure of whole-plant N uptake so that the measured reductant cost of NO₃ assimilation, _Nred (unit of mol e mol¹N), was calculated as:

(4)

where X denotes the roots, stem + petioles or leaves; 16 and 8 refers to hours of the light and dark period, respectively, during a day.

Proportion of Plant NO₃ Assimilation Occurring in Shoots

Two methods were used to calculate the proportion of whole-plant NO₃ assimilation that occurred in the shoot using ratios that could be measured using the methodologies described in this paper. The first assumed that the measured DRUR_Nred values for roots and shoots were directly proportional to the rates of NO₃ assimilation in those plant organs and therefore the ratio of DRUR_Nred^Sht to DRUR_Nred^Rt was related to the proportion of whole-plant NO₃ assimilation that occurred in the shoot as follows:

(5a)

The second incorporated measurements of DRUR_Nred^Rt (which are relatively easy to obtain) and whole-plant ¹⁵N assimilation measurements (a direct measure of N_Red^Plant) to calculate the proportion of whole-plant NO₃ assimilation that occurred in the shoot as follows:

(5b)

where _Nred is the reductant cost of NO₃ assimilation (units of e per NO₃ assimilated).