Inhibition of Nitrate Uptake by Ammonium in Barley. Analysis of Component Fluxes

Abstract

NO₃⁻ uptake by plant roots is rapidly inhibited by exposure to NH₄⁺. The rapidity of the effect has led to the presumption that the inhibition results from the direct effects of NH₄⁺ at the plasma membrane. The mechanism of this inhibition, however, has been in contention. In the present study we used the radiotracer ¹³N to determine the relative effects of short-term exposures to NH₄⁺ on the ¹³NO₃⁻ influx, efflux, and partitioning of absorbed ¹³N in barley (Hordeum vulgare) roots. Plants were grown without NO₃⁻ or NO₂⁻ (uninduced for NO₃⁻ uptake), or with 0.1, 1.0, 10 mm NO₃⁻, or 0.1 mm NO₂⁻ (to generate plant roots induced for NO₃⁻ uptake). Exposure to 1 mm NH₄⁺ strongly reduced influx; the effect was most pronounced in plants induced for NO₃⁻ uptake when NO₃⁻ absorption was measured at low external NO₃⁻. At higher [NO₃⁻] and in uninduced plants the inhibitory effect was much diminished, indicating that NH₄⁺ inhibition of influx was mediated via effects on the inducible high-affinity transport system rather than on the constitutive high-affinity transport system or the low-affinity transport system. Exposure to NH₄⁺ also caused increased NO₃⁻ efflux; the largest effect was at low external [NO₃⁻] in uninduced plants. In absolute terms, the reduction of influx made the dominant contribution to the observed reduction of net uptake of NO₃⁻. Differences in response between plants induced with NO₃⁻ and those induced with NO₂⁻ indicate that NO₂⁻ may not be an appropriate analog for NO₃⁻ under all conditions.

The inhibitory effects exerted by the NH₄⁺ ion upon NO₃⁻ uptake by the roots of higher plants have been studied extensively (Weissman, 1950; Lycklama, 1963; Fried et al., 1965; Minotti et al., 1969; Jackson et al., 1976; Rao and Rains, 1976; Doddema and Telkamp, 1979; MacKown et al., 1982a; Deane-Drummond and Glass, 1983; Rufty et al., 1983; Breteler and Siegerist, 1984; Glass et al., 1985; Ingemarsson et al., 1987; Oscarson et al., 1987; Lee and Drew, 1989; Warner and Huffaker, 1989; de la Haba et al., 1990; Ayling, 1993; Aslam et al., 1994, 1997; Chaillou et al., 1994). It is evident that there are short-term effects of NH₄⁺ on NO₃⁻ uptake that are presumed to result from the direct effects of NH₄⁺ on the plasma membrane; these short-term effects are apparent within minutes of exposure to NH₄⁺. Moreover, longer-term effects due to NH₄⁺ and/or assimilation products of NH₄⁺ are thought to operate at the transcriptional level (Glass and Siddiqi, 1995; Krapp et al., 1998; Zhuo et al., 1999).

Despite the efforts of many investigators, a lack of consensus persists concerning the mechanism(s) responsible for the short-term inhibition of NO₃⁻ uptake by NH₄⁺; specifically, whether the NH₄⁺ effect is achieved by the direct inhibition of influx or by stimulating efflux. Although early reports suggested that NH₄⁺ enhanced NO₃⁻ efflux (Jackson et al., 1976; Doddema and Telkamp, 1979; MacKown et al., 1982a; Deane-Drummond and Glass, 1983; Deane-Drummond, 1985, 1986), later studies using ¹³NO₃⁻ clearly documented an inhibition of influx (Glass et al., 1985; Lee and Clarkson, 1986; Ingemarsson et al., 1987; Oscarson et al., 1987; Lee and Drew, 1989; Ayling, 1993; King et al., 1993).

The debate has recently been revived by Aslam and coworkers (1994, 1997), who concluded that the main effect of NH₄⁺ on net NO₃⁻ uptake was through stimulation of NO₃⁻ efflux; they discounted the significance of influx inhibition. A resolution of this controversy has proved difficult because the experiments have used different species or cultivars; more importantly, different techniques were used to determine NO₃⁻ fluxes. Furthermore, in none of the above studies were both influx and efflux of NO₃⁻ measured directly and simultaneously; rather, conclusions were based upon measurements of net flux or influx or, where efflux was determined, upon the use of NO₃⁻ analogs such as ClO₃⁻ (Deane-Drummond and Glass, 1983; Deane-Drummond, 1985, 1986) and NO₂⁻ (Aslam et al., 1994). Glass et al. (1985) and Siddiqi et al. (1992) have questioned whether analogs of this sort are appropriate. In addition, Glass et al. (1985) and Ingemarsson et al. (1987) have argued that design features of experiments may have caused perturbations from a steady state, which might explain the sometimes large increases in NO₃⁻ efflux that were observed in some of the above studies.

We have designed the present study to address such issues. To eliminate problems associated with the choice of plant material, we used the same barley cultivar (CM-72) used by Aslam and coworkers (1994, 1997), whose studies led to the conclusion that the short-term inhibition of NO₃⁻ uptake by NH₄⁺ resulted exclusively from the stimulation of efflux. In this study we have determined the influx and efflux of NO₃⁻ by using ¹³NO₃⁻ under both steady-state and perturbation protocols, by directly measuring influx and efflux, and by estimating influx calculated from CAE. To investigate possible differential effects of NH₄⁺ on the CHATS, IHATS, and LATS for NO₃⁻ transport, we measured NO₃⁻ uptake at [NO₃⁻] that characterize these transport systems and we used plants induced or uninduced for NO₃⁻ transport. In addition, the experiments were designed to compare the effect of NH₄⁺ on NO₃⁻ influx and efflux in plants induced for NO₃⁻ uptake by prior exposure to either NO₃⁻ or NO₂⁻.

MATERIALS AND METHODS

Plant Growth Conditions

Barley (Hordeum vulgare L. cv CM-72) seeds were surface-sterilized in 1% NaOCl for 10 min, rinsed with deionized water, and germinated in sterilized moist sand in the dark as described by Siddiqi et al. (1989). Seeds were placed on plastic mesh fitted into Plexiglas (Atohaas Americas, Philadelphia, PA) disks, with 40 to 50 seeds per disk for influx experiments and 15 to 20 seeds per disk for efflux experiments (Siddiqi et al., 1989, 1991). After 3 d of germination in the dark, seedlings were transferred to 8-L hydroponic Plexiglas tanks located in walk-in controlled-environment growth chambers. The seedlings grew in hydroponic tanks for 4 d, after which we performed labeling experiments as described below. Growth chambers were maintained at 20°C ± 2°C, 70% RH, and set to a 16-h light/8-h dark photoperiod. Fluorescent lamps (model 1500, F96T12/CW/VHO, 215 W, Philips, Eindhoven, The Netherlands) provided a photon flux of approximately 300 μmol m⁻² s⁻¹, measured at plant level (LI-189 light meter and LI-190SA quantum sensor, LI-COR).

Nutrient Solutions

After the 3-d germination treatment in sand, seedlings were cultivated for 4 d in hydroponic media in 8-L Plexiglas tanks. We used deionized distilled water and reagent-grade chemicals in the preparation of all nutrient solutions. Modified, one-quarter-strength Johnson's nutrient solution (2 mm KH₂PO₄, 2 mm K₂SO₄, 1 mm MgSO₄, 4 mm Ca²⁺ provided as CaSO₄ and/or Ca[NO₃]₂, and the micronutrients 50 μm Cl, 25 μm B, 20 μm Fe as Fe-EDTA, 2 μm Mn, 2 μm Zn, and 0.5 μm Cu) was used in all experiments (Siddiqi et al., 1989). NO₃⁻ was provided (in the form of CA[NO₃]₂) at 0.1, 1.0, or 10 mm starting 24 h before initiating the experiments. When experiments used NO₂⁻ to induce NO₃⁻ transport, it was provided as NaNO₂ (at 0.1 mm). During labeling experiments NH₄⁺ was added as (NH₄)₂SO₄ at 1 mm. Electric circulating pumps (model IC-2, Brinkmann) continuously mixed the nutrient solutions in tanks. Continuous infusion of nutrient stock solution via peristaltic pumps (Technicon Proportioning Pump II, Technicon Instrument Co., Tarrytown, NY) allowed steady-state control of nutrient concentrations in the tanks. We checked the solutions daily for [K⁺] using a spectrophotometer (model 443, Instrumentation Laboratory, Lexington, MA). Powdered CaCO₃ maintained the solution pH at 6.5 ± 0.3. We monitored the pH daily with a microprocessor-based pocket-size pH meter (pH Testr2 model 59000-20, Cole Parmer, Chicago, IL). The [NO₃⁻]_o was measured spectrophotometrically by the method of Cawse (1967).

Influx Analysis

The radiotracer ¹³N (with a half-life of 9.98 min) was produced by the Tri-University Meson Facility cyclotron at the University of British Columbia (Vancouver, Canada) by proton irradiation of water, producing mostly ¹³NO₃⁻ with high radiochemical purity (Kronzucker et al., 1995b). The irradiated solutions were supplied in sealed 20-mL glass vials with a starting activity of 700 to 740 MBq. At this activity level, sufficient counts were present in eluates and plant samples even after loading periods of up to 60 min and a total elution period of 22 min in efflux experiments (see below). Procedures for the removal of radiocontaminants were carried out as described in detail elsewhere (Kronzucker et al., 1995a, 1995b). A volume of 100 mL of purified ¹³NO₃⁻-containing “stock” solution was prepared in a fume hood and transferred into the controlled-environment chambers where the experiments were performed. All uptake solutions were premixed and contained in individual 500-mL plastic vessels behind lead shielding.

The chemical composition of the uptake, prewash, and desorption solutions was identical to the growth solution in the hydroponic tanks (see above) and contained 0.1, 1.0 or 10 mm NO₃⁻. When NH₄⁺ was present in uptake solutions it was provided at a concentration of 1 mm. In experiments where NO₂⁻ was used to induce NO₃⁻ transport (King et al., 1992; Aslam et al., 1997), NO₂⁻ was not present during loading with ¹³NO₃⁻, but only during the induction period (24 h); it was replaced by NO₃⁻ during ¹³NO₃⁻ loading and flux measurement. Uninduced plants received no N during growth but were exposed to 0.1 mm NO₃⁻ for flux determinations.

To minimize plant perturbation during experiments, a syringe was used to add tracer to the individual uptake vessels. At the start of the influx experiments, barley seedlings were transferred from the hydroponic growth tanks to prewash solutions in 1-L vessels for 5 min prior to addition of radioisotope to the uptake solutions. This protocol minimized plant perturbation and allowed the roots to equilibrate to the exact solution temperature and composition used during flux analysis. The roots were then exposed to tracer for 5 min. Immediately after loading with isotope, roots were dipped into nonlabeled solutions for 5 s to minimize the carryover of label by the root surface to the desorption solution. Roots were then desorbed for 2 min in unlabeled solution, which was otherwise chemically identical to the influx solution, to remove the ¹³NO₃⁻ contained in the cell-wall free space. The duration of these steps was based on the half-lives of exchange of NO₃⁻ for the root surface, the free space, and the cytoplasm as determined by efflux analysis (see below; Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b, 1995e).

We chose exposure times of 5 min for ¹³NO₃⁻ influx, because we expected the contribution of tracer efflux from the cytoplasm to be negligible during this time (Lee and Clarkson, 1986; Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b, 1995d, and 1995e). After desorption, seedling roots were excised from the shoots; the roots were spun in a low-speed centrifuge for 30 s to remove any surface liquid; and the fresh weights of the roots and shoots were measured. The plant organs were then introduced into 20-mL scintillation vials, and a γ-counter (Minaxi δ, Auto-γ 5000, Packard Instruments, Meriden, CT) determined the radioactivities of the roots and shoots, measuring the 511-keV positron-electron annihilation radiation generated by the recombination of ambient electrons and β⁺ particles emitted from ¹³N. Using the specific activity (¹³N/[¹³N + ¹⁴N] cpm μmol⁻¹) of the loading solution and the total root fresh weight of each seedling, we calculated the NO₃⁻ fluxes and expressed them in micromoles per gram root fresh weight per hour.

In addition to direct influx determinations by ¹³N count accumulation over the 5-min loading periods (designated as Φ_oc*), influx was also determined by CAE (Φ_oc) and net flux was determined by ¹⁴N depletion over a period of up to 2 h (Φ_net*). We repeated all experiments at least three times. Each experimental treatment consisted of three to four replicates (n ≥ 9).

CAE

The protocol for CAE was essentially as described elsewhere (Lee and Clarkson, 1986; Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b, 1995e). Roots of intact barley seedlings were immersed for 60 min in 120-mL darkened plastic beakers containing the ¹³NO₃⁻-labeled solution. NO₃⁻ concentrations were 0.1, 1.0, or 10 mm. NH₄⁺ was added at a 1 mm concentration unless otherwise indicated for the duration of loading and elution; or it was added only at a specified time during the elution of tracer from the cytoplasm (Figs. 2 and 3) to study the immediate effect of NH₄⁺ upon NO₃⁻ efflux. Pretreatment of uninduced and NO₂⁻-induced plants took place as described above. Conditions closely approximating a steady state with respect to all other nutrients were maintained throughout growth by completely replacing solutions in the 8-L tanks every day.

Figure 2.

¹³NO₃⁻-efflux plot for intact seedlings of cv CM-72 maintained at 0.1 mm [NO₃⁻]_o with a one-time addition (and continued presence) of 1.0 mm NH₄⁺ during cytoplasmic efflux (at 12 min during tracer elution). Inset shows magnified cytoplasmic exchange.

Figure 3.

¹³NO₃⁻-efflux plot for intact seedlings of cv CM-72 maintained at 0.1 mm [NO₃⁻]_o with two-time addition and subsequent withdrawal of 1.0 mm NH₄⁺ during cytoplasmic efflux (additions at 5 and 16 min; withdrawals at 12 and 21 min during tracer elution). Inset shows magnified cytoplasmic exchange.

We maintained steady-state conditions during loading and elution. We chose the duration of the loading period on the basis of the half-lives of exchange for the cytoplasmic compartment for NO₃⁻ in barley (compare below with Siddiqi et al., 1991). Therefore, 60 min of exposure to tracer ensured that cytoplasmic specific activity was ≥95% of that in the loading solution (Kronzucker et al., 1995e). After loading with ¹³N, seedlings were transferred to efflux funnels (Siddiqi et al., 1991; Kronzucker et al., 1995b) and the roots were eluted with 20-mL aliquots of nonradioactive solution after varying time intervals. Using an elution protocol lasting 22 min, these time intervals ranged from 5 s to 2 min, as described by Kronzucker et al. (1995b), except when we monitored the response of the NO₃⁻ efflux to the NH₄⁺ addition during elution (Figs. 2 and 3); we used 1-min intervals in those cases to ensure appropriate time resolution.

Eluates from a total of 25 time intervals were collected, and the γ-counter (see above) determined the radioactivities of 20-mL subsamples from each eluate. After each final elution, we excised the seedling roots from the shoots, spun the roots in a low-speed centrifuge for 30 s to remove surface liquid, and determined the fresh weights of the roots and shoots. We then introduced the plant organs into 20-mL scintillation vials and determined the radioactivities of the roots and shoots as described previously for the influx experiments. We repeated the experiments three times with two replicates per experiment. We pooled the data from several experiments (n ≥ 6) to calculate means and se. Symbols and calculation of fluxes in CAE were as follows: Φ_co, efflux from the cytoplasmic compartment at time 0 divided by the specific activity of ¹³N in the loading solution; Φ_net, net flux, obtained from the accumulation of ¹³N in the plants at the end of the loading period (60 min); Φ_oc, unidirectional influx, calculated from Φ_net + Φ_co; Φ_xylem, flux of ¹³N to the shoot at the end of the elution period; and Φ_red/vac, combined flux to reduced N and the vacuole, resulting in Φ_net − Φ_xylem. Calculations of half-lives of exchange and cytosolic concentrations were done as described in detail elsewhere (Lee and Clarkson, 1986; Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b, 1995c, 1995e).

RESULTS AND DISCUSSION

Effect of NH₄⁺ on Half-Lives of Cellular NO₃⁻ Exchange

As in our previous studies with barley, rice, and spruce (Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b, 1997; H.J. Kronzucker, A.D.M. Glass, and M.Y. Siddiqi, unpublished results), CAE revealed NO₃⁻ exchange with three subcellular compartments (Fig. 1). These corresponded to a surface film, a binding component in the cell wall, and the cytoplasm, an interpretation substantiated by previously reported compartment identity tests for the CAE technique with ¹³N (Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b, 1995e). Half-lives of exchange for the surface film and the cell wall free space were very similar (approximately 2 and 30 s, respectively) to those reported in our previous studies (Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b) and did not change significantly as a function of [NO₃⁻]_o. The half-lives for cytoplasmic NO₃⁻ exchange without NH₄⁺ additions are shown in Table I. Cytoplasmic half-life values of NO₃⁻ exchange ranged from 7.75 to 12.94 min, with slightly shorter half-lives at higher [NO₃⁻]_o and longer half-lives in NO₂⁻-induced plants (Lee and Clarkson, 1986; Siddiqi et al., 1991; Devienne et al., 1994; Kronzucker et al., 1995a, 1995b, 1995c, 1995e, 1997).

Figure 1.

Representative semilogarithmic plots for the rate of release of ¹³NO₃⁻ versus time of elution for roots of intact cv CM-72 seedlings maintained under steady-state conditions of 0.1 mm [NO₃⁻]_o without NH₄⁺ (•) and following the addition of 1 mm NH₄⁺ (□). Plots include linear regression lines for the three phases of efflux: I, surface film; II, cell wall; and III, cytoplasm. Regression lines are dashed for the plus-NH₄⁺ treatment and solid for the control (phases I and II overlapped). For derivation of compartmental parameters, see text.

Table I.

Half-lives of NO₃⁻ exchange for the cytoplasmic compartment and cytoplasmic [NO₃⁻] ([NO₃⁻]_cyt) in roots of intact cv CM-72 plants, determined by compartmental analysis

Treatment	t1/2, Cytoplasm		[NO3−]cyt
	−	+	−	+
	min		mm
0.1 mm NO3−	10.94  ± 0.57	14.78  ± 1.04	41  ± 2.89	38.75  ± 3.06
1.0 mm NO3−	8.52  ± 0.4	13.32  ± 0.99	51.69  ± 1.49	59.4  ± 2.8
10 mm NO3−	7.75  ± 0.37	6.75  ± 0.24	76.18  ± 1.03	67.27  ± 9.34
Uninduced	9.17  ± 0.98	8.55  ± 0.49	2.05  ± 0.45	2.19  ± 0.31
NO2− induced	12.94  ± 0.53	17.59  ± 0.98	48.8  ± 3.1	50.6  ± 2.71

Plants were exposed to and labeled at the indicated concentrations of NO₃⁻ (steady state). Uninduced plants were grown in N-free solution but were exposed to 0.1 mm NO₃⁻ during labeling and elution. NO₂⁻-induced plants were exposed to 0.1 mm NO₂⁻ for 24 h prior to labeling and elution at 0.1 mm NO₃⁻. NH₄⁺ was present at 1.0 mm during labeling and elution in + treatments. Data are ±se (n ≥ 6).

In general, half-life values for the cytoplasm were longer in the cv CM-72 used in the present study compared with the Klondike variety of barley used in previous studies (Siddiqi et al., 1991; Devienne et al., 1994; Kronzucker et al., 1995a, 1995b), indicating a larger relative accumulation capacity for NO₃⁻ (at comparable fluxes) in the former variety. The addition of NH₄⁺ significantly affected cytoplasmic exchange kinetics in NO₂⁻-induced plants as well as in plants grown at 0.1 and at 1 mm [NO₃⁻]_o: Half-lives increased in all of these cases (Fig. 1), whereas they remained unaffected in uninduced plants and at 10 mm [NO₃⁻]_o (Table I). These differences in half-life values are direct reflections of differences in flux partitioning, as discussed below.

Effect of NH₄⁺ on NO₃⁻ Uptake and Subcellular N-Flux Partitioning

The addition of NH₄⁺ led to a reduction of net NO₃⁻ uptake, as estimated by compartmental analysis (Table II) or as ascertained by direct methods (Table III). Moreover, there was a close correspondence between the values for the percentage of inhibition determined by CAE (Table II) and those determined independently by ¹⁴N depletion (Table III), although the absolute flux values tended to be higher in CAE determinations. The inhibitory effect of externally applied NH₄⁺ is in agreement with other studies (see the introduction), although genetic variability and even a stimulation of NO₃⁻ uptake by NH₄⁺ in isolated cases (Bloom and Finazzo, 1986) have been reported in the responses, which ranged from strong to low levels of inhibition (Schrader et al., 1972; Pan et al., 1985). In the present study the inhibition of net NO₃⁻ uptake by 1.0 mm NH₄⁺ depended upon [NO₃⁻] and the N species used to induce NO₃⁻ uptake prior to exposure to NH₄⁺. In plants that were uninduced for NO₃⁻ uptake, NH₄⁺ reduced net NO₃⁻ uptake by approximately 25% when measured in solutions containing 0.1 mm [NO₃⁻]_o. In plants induced for NO₃⁻ uptake, the corresponding values for inhibition were 35% and 25% when net uptake was measured in solutions containing 0.1 and 1.0 mm [NO₃⁻]_o, respectively, whereas at 10 mm [NO₃⁻]_o, 1 mm external NH₄⁺ had no significant effect. The inhibition of net NO₃⁻ uptake was significantly less (approximately 14%–17%) when plants were induced with NO₂⁻ and NO₃⁻ uptake was measured using 0.1 mm [NO₃⁻]_o. The shortcomings of measuring net fluxes by depletion have been discussed previously (Kronzucker et al., 1995d, 1996). In particular, unless short-term (5–10 min) estimates are used, it is possible that plant acclimation will occur in response to declining [NO₃⁻]_o during the measurement.

Table II.

Component fluxes of NO₃⁻ as determined by compartmental analysis

Treatment	Φoc		Φco		Φnet		Φred/vac		Φ*xylem
	−	+	−	+	−	+	−	+	−	+
	μmol g−1 h−1
0.1 mm NO3−	8.61  ± 0.77	5.77  ± 0.59	0.57  ± 0.02	0.84  ± 0.13	7.57  ± 0.76	4.93  ± 0.7	5.3  ± 0.6	3.49  ± 0.62	2.27  ± 0.25	1.44  ± 0.1
1.0 mm NO3−	12.93  ± 0.88	10.02  ± 0.13	1.6  ± 0.56	1.46  ± 0.36	11.33  ± 1.44	8.55  ± 1.06	6.85  ± 1.22	5.86  ± 1.08	4.47  ± 0.22	2.69  ± 0.02
10 mm NO3−	20.94  ± 0.73	21.58  ± 0.57	10.56  ± 3.97	12.14  ± 0.98	10.38  ± 4.7	9.44  ± 1.54	6.68  ± 2.08	5.95  ± 1.76	3.7  ± 2.62	3.49  ± 2.17
Uninduced	0.53  ± 0.12	0.48  ± 0.04	0.07  ± 0.01	0.13  ± 0.01	0.46  ± 0.1	0.34  ± 0.02	0.36  ± 0.07	0.28  ± 0.03	0.096  ± 0.03	0.055  ± 0.005
NO2− induced	8.1  ± 0.69	7.16  ± 0.74	0.29  ± 0.02	0.47  ± 0.09	7.8  ± 0.75	6.69  ± 0.49	6.67  ± 0.62	5.46  ± 0.49	1.13  ± 0.22	1.23  ± 0.09

Barley plants were exposed to and labeled at the indicated concentrations of NO₃⁻ for 24 h prior to experiments. Uninduced plants were grown in N-free solution but exposed to 0.1 mm NO₃⁻ during labeling and elution. “NO₂⁻-induced” plants were exposed to 0.1 mm NO₂⁻ for 24 h prior to labeling and elution at 0.1 mm NO₃⁻. NH₄⁺ was present at 1.0 mm during labeling and elution in + treatments. Data are ±se (n ≥ 6).

Table III.

Estimates of NO₃⁻ influx and net flux into roots of intact cv CM-72 plants by methods independent of compartmental analysis

Treatment	Φoc		Φnet
	−	+	−	+
	μmol g−1 h−1
0.1 mm NO3−	5.16  ± 0.45	3.26  ± 0.47	3.97  ± 0.7	2.56  ± 0.61
1 mm NO3−	6.91  ± 0.34	5.37  ± 0.38	8.57  ± 0.16	6.34  ± 0.27
10 mm NO3−	10.52  ± 0.4	10.04  ± 0.93	N/D	N/D
Uninduced	0.42  ± 0.02	0.24  ± 0.02	N/D	N/D
NO2− induced	4.49  ± 0.16	3.87  ± 0.08	5.55  ± 0.13	4.59  ± 0.16

Plants were exposed to, and fluxes were measured at, the indicated concentrations of NO₃⁻ (steady state). Uninduced plants were grown in N-free solution prior to flux measurement at 0.1 mm NO₃⁻. NO₂⁻-induced plants were exposed to 0.1 mm NO₂⁻ for 24 h prior to flux measurement at 0.1 mm NO₃⁻. NH₄⁺ was present at 1.0 mm during uptake in + treatments. Data are ±se (n ≥ 9).

NH₄⁺ had a distinct and more potent effect on NO₃⁻ uptake in the high-affinity transport range (i.e. on the IHATS), which is evident below 1 mm [NO₃⁻]_o, than in the range of the LATS, which operates at [NO₃⁻]_o ≥ 1 mm (Siddiqi et al., 1989; King et al., 1993; Kronzucker et al., 1995d). This provides additional support for the argument that high- and low-affinity transport systems are biochemically distinct modes of transport (for review, see Glass and Siddiqi, 1995). NO₂⁻-induced plants were substantially less sensitive to NH₄⁺ inhibition of NO₃⁻ uptake than NO₃⁻-induced plants. Aslam and coworkers (1994, 1997) used plants induced by and “labeled” with NO₂⁻ as model systems for NO₃⁻-induced plants. Although NO₂⁻ and NO₃⁻ have been shown to act competitively at the level of uptake (Aslam et al., 1992; Siddiqi et al., 1992), the present findings suggest that NO₂⁻ may not serve as a satisfactory quantitative analog for NO₃⁻. This is consistent with the finding by Siddiqi et al. (1992) that NO₂⁻ was not capable of inducing NO₃⁻ reductase activity in barley. Aslam et al. (1987, 1993, 1997), however, reached the opposite conclusion.

The effect of 1 mm NH₄⁺ on ¹³NO₃⁻ influx followed the same pattern (with one exception) as that observed for net uptake, i.e. the extent of inhibition declined with increasing [NO₃⁻]_o, with 33% inhibition at 0.1 mm, 23% at 1.0 mm, and no effect at 10 mm. Inhibition was much smaller in NO₂⁻-induced plants (11.6%). This was true when influx was determined by CAE and count accumulation after a 5-min exposure to tracer (Tables II and III). The one exception was that influx in uninduced plants (determined by CAE) was unaffected by NH₄⁺, in contrast to the situation for net uptake by uninduced plants. We interpret this result to indicate that, like LATS, the NO₃⁻ influx via CHATS (King et al., 1992; Kronzucker et al., 1995d) is unaffected by NH₄⁺. That exposure to NH₄⁺ diminished net uptake in uninduced plants indicates an effect on efflux of NO₃⁻ in uninduced plants.

We measured CAE on uninduced plants by exposing roots to 0.1 mm [NO₃⁻]_o for the duration of the efflux analyses (the labeling and elution procedures lasted 82 min); by this time, the physiological induction of the IHATS and NO₃⁻ reductase would be relatively small (Friemann et al., 1992; Glass and Siddiqi, 1995; Kronzucker et al., 1995a, 1995b). In contrast to its lack of effect on constitutive influx, NH₄⁺ stimulated NO₃⁻ efflux by as much as 86% (Table II) in uninduced plants. Although not measured directly, efflux was probably even larger during the shorter 5-min exposures to NH₄⁺ (see discussion of short-term efflux enhancements below), which may explain the apparent depression of influx that we saw in the short-term determinations after the addition of NH₄⁺ (Table III), but not in the CAE determinations (Table II).

Efflux stimulation was also substantial in NO₂⁻-induced plants (approximately 62%), whereas in plants grown at 0.1 mm [NO₃⁻]_o, efflux was enhanced by less than 50% and no efflux stimulation at all was seen in plants grown at 1.0 and 10 mm [NO₃⁻]_o (Table II; Fig. 1). NO₃⁻ efflux (expressed as a percentage of influx) also increased as [NO₃⁻]_o was raised from 0.1 to 10 mm even in the absence of NH₄⁺ (Table II), as previously reported (Siddiqi et al., 1991; Kronzucker et al., 1995b; Volk, 1997). Our results confirm a distinct difference between inducible high-affinity transport and low-affinity transport; they also confirm the fact that NO₂⁻ induction of NO₃⁻ transport cannot be used as a quantitative model for NO₃⁻ induction and provision under steady-state conditions. The latter point is particularly important, because experiments on plants pretreated in this way have led to the conclusion that the NH₄⁺ inhibition of net NO₃⁻ uptake results exclusively from the effects on NO₃⁻ efflux and that influx is unaffected (Aslam et al., 1994). Furthermore, the relative effect of NH₄⁺ was high in the present study (>50% stimulation) because efflux is typically low under control conditions, and the absolute contribution to reduced net uptake was still small in comparison with the contribution arising from reduced influx.

Our experiments show that in NO₂⁻-induced plants, the combined flux of ¹³NO₃⁻ and ¹³N-assimilation products to the shoot (Φ*_xylem) was unaffected by the addition of NH₄⁺, whereas it was reduced in all other treatments (Table II). Also, Φ*_xylem in NO₂⁻-induced plants was substantially lower than in NO₃⁻-induced plants, and approximated that of plants induced at 0.1 mm [NO₃⁻]_o after the application of 1 mm NH₄⁺. Because significant suppression of NO₃⁻ reductase activity by NH₄⁺ is well documented (MacKown et al., 1982a, 1982b; Breteler and Siegerist, 1984; Pan et al., 1985; de la Haba et al., 1990; Aslam et al., 1997; however, see Oaks et al., 1979), Φ*_xylem under these conditions will be mostly in the form of ¹³NO₃⁻ after NH₄⁺ addition. Therefore, the lack of an effect of NH₄⁺ on Φ*_xylem in NO₂⁻-induced plants (measured at 0.1 mm [NO₃⁻]_o) appears to support the biochemically based conclusion arrived at by Siddiqi et al. (1992) and King et al. (1993) that NO₃⁻ reductase activity is not induced to any significant extent by NO₂⁻ in barley roots, in contrast to its full induction by NO₃⁻. Claims to the contrary have been made by Aslam et al. (1987, 1993; see Glass and Siddiqi, 1995).

Our CAE analyses do not allow a separation of the reduced and unreduced components of the xylary ¹³N-translocation term, nor do they permit separation of the components of Φ_red/vac, namely the biochemical N flux to reduced N, and the N flux to the vacuole (Lee and Clarkson, 1986; Siddiqi et al., 1991; Kronzucker et al., 1995a, 1995b). However, given that the cytoplasmic [NO₃⁻] values in NO₂⁻- and NO₃⁻-induced plants were indistinguishable after ¹³N loading (Table I), and thus similar values of Φ*_xylem for NO₃⁻ are to be expected, we must conclude that NO₂⁻ induction leads to a relatively greater stimulation of the NO₃⁻ flux to the vacuole (in both the control and NH₄⁺-treated plants). The reduction in Φ_red/vac after NH₄⁺ addition to NO₂⁻-induced plants (Table II) appears to be a direct result of the inhibition of influx and the stimulation of efflux rather than the effect upon NO₃⁻ reductase activity.

The clear differences in flux partitioning between NO₃⁻- and NO₂⁻-induced plants revealed by CAE lead us to caution against the use of NO₂⁻ as an “analog” for NO₃⁻. Similar concerns regarding the use of ³⁶ClO₃⁻ as an analog for NO₃⁻ were expressed in earlier work (Deane-Drummond and Glass, 1983; Dean-Drummond, 1985, 1986), although the basis of the failure to faithfully trace NO₃⁻ in the case of ClO₃⁻ was more straightforward (Glass et al., 1985; Lee and Drew, 1989; Siddiqi et al., 1992; Aslam et al., 1994; Glass and Siddiqi, 1995). Doddema and Telkamp (1979) also observed a significant rise in NO₃⁻ efflux upon the addition of NH₄⁺; however, this response was transient and restricted to the perturbational condition (see below). Our present analyses demonstrate that an enhancement of the efflux component makes only a small contribution to the reduction of net uptake, whereas the principal effect of NH₄⁺ on NO₃⁻ uptake comes through the inhibition of influx (except in uninduced plants and in plants pretreated with NO₂⁻, where the efflux contribution is magnified).

Kinetics of the Response

It has been shown previously that the inhibition of NO₃⁻ influx by NH₄⁺ is an immediate phenomenon, detectable even within 15 s (Glass et al., 1985; Ingemarsson et al., 1987; Lee and Drew, 1989; Ayling, 1993; Aslam et al., 1994), and that it is reversible with relaxation times of only 2 to 3 min (Lee and Drew, 1989). In the present study we tested the immediacy and reversibility of the stimulation of NO₃⁻ efflux by NH₄⁺ (in plants grown with 0.1 mm [NO₃⁻]_o); we added NH₄⁺ under perturbational conditions only during the elution of the cytoplasmic compartment in ¹³NO₃⁻ efflux experiments. Figures 2 and 3 show that the effect of NH₄⁺ on NO₃⁻ efflux was evident immediately after its addition to the elution solutions (Fig. 2). Upon withdrawal of NH₄⁺ from these solutions, NO₃⁻ efflux rapidly returned to normal (Fig. 3); repeated additions/withdrawals confirmed that the process was fully reversible. Notwithstanding this clear response, the absolute contribution to the inhibition of NO₃⁻ net uptake was small compared with the large reduction of influx. Moreover, in a separate study on rice (H.J. Kronzucker, A.D.M. Glass, and M.Y. Siddiqi, unpublished results), we found that in long-term studies under steady-state conditions in which both NO₃⁻ and NH₄⁺ are provided, NO₃⁻ efflux was reduced rather than enhanced. Under these conditions efflux as a percentage of influx was very similar to that seen when only NO₃⁻ was provided; thus the efflux enhancement appears to be temporary.

The rapidity of the response to NH₄⁺ on both the influx and efflux components of NO₃⁻ uptake provides a compelling argument that the NH₄⁺ effect occurs directly at the plasma membrane. Lee and Drew (1989) demonstrated a logarithmic relationship between the inhibition of NO₃⁻ influx by NH₄⁺ and external [NH₄⁺], which led to the suggestion that membrane depolarization by NH₄⁺ may inhibit the NO₃⁻/2H⁺ cotransport system due to effects on the proton motive force (Ullrich et al., 1984; Ayling, 1993). However, the provision of K⁺, which also depolarizes the plasma membrane to an extent similar to that of NH₄⁺, fails to inhibit NO₃⁻ uptake (Glass and Siddiqi, 1995; Wang et al., 1996), arguing for a more specific effect of NH₄⁺. Given the rapidity of the response, the inhibition probably occurs allosterically, rather than by involving the products of NO₃⁻ reduction and N assimilation, or possibly the effects of transcription or translation.

Similar conclusions have been reached by others (Deane-Drummond and Glass, 1983; Ingemarsson et al., 1987; Lee and Drew, 1989; Warner and Huffaker, 1989; Aslam et al., 1994). In agreement with de la Haba et al. (1990) and Aslam et al. (1994), we found that pretreatment of barley and rice plants with the Gln synthetase inhibitor Met sulfoximine for 6 h at 1 mm did not alleviate the inhibitory effect exerted by externally added NH₄⁺ (data not shown). Thus it seems unlikely that N assimilates downstream of NH₄⁺ are involved in the inhibition of NO₃⁻ uptake. We must stress that these conclusions apply only to the rapid effects of NH₄⁺ on NO₃⁻ uptake; some (Krapp et al., 1998; Zhuo et al., 1999) have suggested that there may be long-term effects of Gln and other amino acids at the transcription level.

Summary

Our analyses provide evidence that the inhibitory effect of NH₄⁺ upon NO₃⁻ uptake is mediated primarily by inhibiting NO₃⁻ influx, with only a small contribution from the enhancement of NO₃⁻ efflux, which: (a) is both transient and reversible, (b) is associated with a large efflux only in uninduced plants and plants induced by NO₂⁻ (i.e. under conditions where influx is very low), (c) is dependent on [NO₃⁻]_o, (d) is strong for IHATS but small for CHATS and LATS, (e) occurs directly at the plasma membrane (i.e. it does not involve NO₃⁻ reduction or N-assimilation products in the short term, although it may in the long-term); and (f) cannot be modeled quantitatively by the use of NO₂⁻ as an analog of NO₃⁻.

Abbreviations:

CAE

compartmental analysis by efflux

CHATS

constitutive high-affinity transport system

IHATS

inducible high-affinity transport system

LATS

low-affinity transport system

[NO₃⁻]_o

external [NO₃⁻] Φ, ionic (N) flux (component fluxes denoted by subscripts, as indicated in text)