Dual Mechanisms of Nitrate in Alleviating Ammonium Toxicity: Enhanced Photosynthesis and Optimized Ammonium Utilization in Orychophragmus violaceus

Abstract

Ammonium (NH₄⁺) toxicity impairs plant growth, but nitrate (NO₃⁻) can mitigate this effect through unresolved mechanisms. Using leaf δ¹³C values (photosynthetic capacity) and a bidirectional ¹⁵N tracer (NH₄⁺ assimilation efficiency and source utilization), this study investigated these mechanisms in 35-day-old Orychophragmus violaceus plantlets grown in modified Murashige and Skoog media under varying NH₄⁺:NO₃⁻ ratios. ¹⁵N isotope fractionation during NH₄⁺ (same fixed 20 mM NH₄Cl) assimilation decreased with increasing NO₃⁻ supply (10, 20, and 40 mM NaNO₃). Under 20 mM NH₄⁺ (δ¹⁵N = −2.64‰) at two ¹⁵NO₃⁻-labels (δ¹⁵N-NO₃⁻ = 8.08‰, low ¹⁵N, L) and (δ¹⁵N-NO₃⁻ = 22.67‰, high ¹⁵N, H), increasing NO₃⁻ concentrations enhanced NO₃⁻ assimilation, alleviating acidic stress from NH₄⁺ and improving photosynthesis. Higher NO₃⁻ levels also increased NH₄⁺ utilization efficiency, reducing futile NH₄⁺ cycling and decreasing associated ¹⁵N fractionation during assimilation. Our results demonstrate that NO₃⁻ alleviates NH₄⁺ toxicity primarily by enhancing photosynthetic performance and optimizing NH₄⁺ utilization efficiency.

1. Introduction

The major inorganic nitrogen (N) sources utilized by plants are nitrate (NO₃⁻) and ammonium (NH₄⁺). Compared to NO₃⁻, NH₄⁺ is often considered the preferred N source due to its lower energy requirement for assimilation [1,2]. However, when plants are exposed to elevated concentrations of NH₄⁺ in the environment, particularly from sources such as untreated wastewater effluents [3], or when plants are exclusively supplied with NH₄⁺, they commonly exhibit symptoms including leaf chlorosis and growth suppression, a phenomenon known as NH₄⁺ toxicity [4,5,6,7]. Interestingly, numerous studies have demonstrated that even a small amount of NO₃⁻ can effectively alleviate NH₄⁺ toxicity, a process termed NO₃⁻-mediated mitigation of NH₄⁺ toxicity [4,5,8,9]. Nevertheless, inadequate NO₃⁻ supply results in persistent NH₄⁺ toxicity symptoms [10], suggesting that the mitigation effect depends critically on NO₃⁻ concentration.

Significant progress has been made in understanding the mechanisms underlying NH₄⁺ toxicity. Proposed contributors include futile NH₄⁺ cycling [11,12], rhizosphere acidification [13,14], inhibition of cation uptake [5,15], damage to the photosynthetic system [16,17], and reactive oxygen species (ROS) accumulation [8,18]. Given these established mechanisms, the mitigation of NH₄⁺ toxicity by NO₃⁻ supplementation likely involves counteracting one or more of these detrimental processes.

The N sources used in the Murashige and Skoog (MS) medium [19] comprise NO₃⁻ and NH₄⁺ with a ratio of 2:1 (NO₃⁻:NH₄⁺) and a total N concentration of 60 mM. Although NO₃⁻ is generally not toxic even at high concentrations [20,21], the relatively high NO₃⁻ in the MS medium may lead to inefficient N utilization. A previous study indicates that plantlets preferentially assimilate NH₄⁺ as their primary N source, even when its concentration is only half that of NO₃⁻ [22]. Therefore, while maintaining the NH₄⁺ concentration at 20 mM, optimizing NO₃⁻ levels could enhance N use efficiency and simultaneously provide insights into the mechanism of NO₃⁻-mediated mitigation of NH₄⁺ toxicity.

Sucrose (typically at 3% w/v) is widely used in the MS medium as a carbon (C) and energy source for plantlets [23,24]. While sucrose supports heterotrophic growth, CO₂ enables autotrophic growth [24]. Consequently, the leaf δ¹³C values of plantlets reflects the combined contribution of C from sucrose utilization and CO₂ assimilation. Carbon isotope discrimination differs significantly between these two processes: CO₂ assimilation exhibits ~20‰ discrimination [25], whereas the sucrose utilization shows a much smaller ~2.54‰ discrimination [10]. By measuring the δ¹³C values of both the sucrose source and atmospheric CO₂, the proportion of C assimilated from CO₂ can be estimated using the leaf δ¹³C values. Therefore, leaf δ¹³C serves as an indirect indicator of photosynthetic capacity [24]. Given this relationship, comparing the leaf δ¹³C values under varying NH₄⁺ to NO₃⁻ ratios provides a means to evaluate photosynthetic capacity. This approach offers an insight into how NH₄⁺ toxicity affects plantlet photosynthesis.

Under high NH₄⁺ conditions, plants mitigate NH₄⁺ toxicity either by exporting excess NH₄⁺ or enhancing its assimilation [26,27]. Both processes are energetically demanding, increasing carbohydrate catabolism and decreasing overall plant carbon content. Crucially, NH₄⁺ assimilation through glutamine synthetase (GS) and NO₃⁻ reductase (NR) exhibits distinctive N isotope discrimination (Δ¹⁵N) [28,29]: GS exhibits Δ¹⁵N ≈ 16.8‰ [30], while NR shows stronger discrimination at 25.1‰ [31]. Effluxed, unassimilated NH₄⁺ becomes enriched in ¹⁵N [31], meaning extensive futile NH₄⁺ cycling produces ¹⁵N-depleted assimilates. Thus, the Δ¹⁵N of NH₄⁺-derived assimilates may indicate futile cycling extent. However, quantifying Δ¹⁵N from NH₄⁺ assimilation under mixed N sources remains challenging.

In this study, root formation in plantlets was suppressed by cytokinin and auxin levels in the culture medium, restricting NO₃⁻ and NH₄⁺ assimilation primarily to leaves. Consequently, the leaf δ¹⁵N values reflect the integrated ¹⁵N signatures of both NO₃⁻ and NH₄⁺ sources. Using clonal plantlets grown uniformly minimized individual variability. To disentangle NO₃⁻ and NH₄⁺ assimilation effects, we employed a bidirectional stable N isotope tracer technique [10,22]. Two treatments were applied: L (NO₃⁻-δ¹⁵N = 8.08‰) and H (NO₃⁻-δ¹⁵N = 22.67‰), differing solely in NO₃⁻ isotopic composition. This approach enables quantification of (1) the Δ¹⁵N associated with NH₄⁺ assimilation and (2) the relative contributions of NO₃⁻ and NH₄⁺ to N uptake.

In this study, we employed bidirectional ¹⁵N labeling and gas exchange-based carbon isotope measurements to examine Orychophragmus violaceus (Brassicaceae) plantlets grown under different inorganic N regimes. Orychophragmus violaceus is a plant adapted to karst environments, exhibiting a robust capacity for nitrate assimilation [22]. The NH₄⁺ concentration was maintained at 20 mM across all treatments. The objectives were three-fold: (1) to quantify the Δ¹⁵N values associated with NH₄⁺ assimilation and to determine the relative contributions of NO₃⁻ and NH₄⁺ to total N uptake under varying NO₃⁻:NH₄⁺ ratios; (2) to assess the photosynthetic performance of plantlets exposed to different inorganic N source ratios; and (3) to elucidate the mechanisms underlying NO₃⁻-mediated mitigation of NH₄⁺ toxicity. The expected results will help resolve conflicting reports in the literature regarding NO₃⁻’s role in NH₄⁺ detoxification.

2. Materials and Methods

2.1. Plant Materials and Experimental Design

The experiment was conducted in November 2016. All reagents were analytical grade. Sodium nitrate (δ¹⁵N-NO₃⁻ = 8.08‰), potassium chloride, magnesium sulfate heptahydrate, potassium phosphate monobasic, calcium chloride dihydrate, and sucrose were provided by Jinshan Chemical Reagent Co., Ltd. (Chengdu, China). Sodium nitrate (δ¹⁵N-NO₃⁻ = 22.67‰) was provided by Kemiou Chemical Reagent Co., Ltd. (Tianjin, China). Ammonium chloride (δ¹⁵N = −2.64‰) was provided by Dengke Chemical Reagent Co., Ltd. (Tianjin, China). Manganese sulfate monohydrate, zinc sulphate heptahydrate, boric acid, potassium iodide, sodium molybdate dihydrate, copper sulfate pentahydrate, cobalt chloride hexahydrate, ferrous sulfate heptahydrate, and ethylenediaminetetraacetic acid disodium salt dihydrate were provided by Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). Inositol, nicotinic acid, vitamin B1, vitamin B6, glycine, agar, α-naphthylacetic acid, and 6-benzylaminopurine were provided by Solarbio Technology Co., Ltd. (Beijing, China). The seeds of O. violaceus were collected from the garden of the Institute of Geochemistry, Chinese Academy of Sciences (Nanming District, Guiyang City, Guizhou Province, China). Individual shoots of O. violaceus plantlets (mean fresh weight = 0.13 g) were cultured as explants in a modified Murashige and Skoog medium [19]. This medium contained 20 mM NH₄Cl (δ¹⁵N = −2.64‰), three NO₃⁻ concentrations (10, 20, and 40 mM NaNO₃), growth regulators as 0.2 mg·L⁻¹ α-naphthylacetic acid and 2.0 mg·L⁻¹ 6-benzylaminopurine, 100 mg·L⁻¹ inositol, 0.5 mg·L⁻¹ nicotinic acid, 0.1 mg·L⁻¹ vitamin B1, 0.5 mg·L⁻¹ vitamin B6, 2 mg·L⁻¹ glycine, 16.9 mg·L⁻¹ MnSO₄·1H₂O, 8.6 mg·L⁻¹ ZnSO₄·7H₂O, 6.2 mg·L⁻¹ H₃BO₃, 0.83 mg·L⁻¹ KI, 0.25 mg·L⁻¹ Na₂MoO₄·2H₂O, 0.025 mg·L⁻¹ CuSO₄·5H₂O, 0.025 mg·L⁻¹ CoCl₂·6H₂O, 27.8 mg·L⁻¹ FeSO₄·7H₂O, 37.3 mg·L⁻¹ C₁₀H₁₄N₂O₈Na₂·2H₂O, 1.4 g·L⁻¹ KCl, 0.37 g·L⁻¹ MgSO₄·7H₂O, 0.17 g·L⁻¹ KH₂PO₄, 0.44 g·L⁻¹ CaCl₂·2H₂O, 30 g·L⁻¹ sucrose, and 7.5 g·L⁻¹ agar. We designed three NO₃⁻:NH₄⁺ ratio treatments, and each ratio was further divided into two ¹⁵N-labeling groups based on NO₃⁻ source: low (L) (δ¹⁵N-NO₃⁻ = 8.08‰) and high (H) (δ¹⁵N-NO₃⁻ = 22.67‰). The medium pH was adjusted to 5.8 before dispensing 50 mL aliquots into Erlenmeyer flasks. The Erlenmeyer flask was sealed with a piece of vented sealing film (dimensions: 12 cm × 12 cm, vented membrane diameter: 3 cm, pore size: 0.2–0.3 μm), thereby enabling gas exchange with the ambient atmosphere. The Erlenmeyer flasks were autoclave-sterilized at 121 °C for 20 min. Plantlets were cultured under controlled conditions at 50 μmol m⁻² s⁻¹ photosynthetic photon flux density (PPFD) and 25 ± 2 °C and a 12 h photoperiod. Root formation was suppressed by the cytokinin/auxin balance, indicating that N assimilation occurred primarily within the shoot tissues.

2.2. Harvest and Biomass Measurements

Following a five-week growth period, O. violaceus plantlets were harvested. Each plantlet was carefully removed from its Erlenmeyer flask. Fresh weight (FW) was recorded immediately, and then dried at 60 °C to constant weight to determine dry weight (DW). Biomass increase was calculated as the difference between the final plantlet FW and the initial shoot FW (0.13 g). Dried leaf samples were finely ground using an agate mortar for subsequent analyses.

2.3. Chlorophyll Concentration Determination

Fresh leaf samples (0.1 g each) were flash-frozen in liquid nitrogen and homogenized using a mortar and pestle. The ground tissue was extracted in 10 mL of 95% ethanol for 24 h at 4 °C in darkness. After centrifugation (4000× g, 10 min, 4 °C), the supernatant absorbance was measured at 665 and 649 nm using spectrophotometer measurements. The chlorophyll (Chl a and Chl b) concentrations were calculated according to Alsaadawi et al. [32] using the following equations:

$$\mathrm{Chl}\mathrm{a}(\mathrm{mg}/\mathrm{g})=(13.70\times {\mathrm{A}}_{665}-5.76\times {\mathrm{A}}_{649})\times \mathrm{V}/1000/\mathrm{W}$$

(1)

$$\mathrm{Chl}\mathrm{b}(\mathrm{mg}/\mathrm{g})=(25.80\times {\mathrm{A}}_{649}-7.60\times {\mathrm{A}}_{665})\times \mathrm{V}/1000/\mathrm{W}$$

(2)

where A₆₆₅ and A₆₄₉ represent the absorbance at 665 and 649 nm, respectively. V is the volume of sample (0.01 L). W is the weight of fresh leaves (0.1 g).

2.4. Analysis of Leaf Nitrogen and Carbon Content

The analysis of leaf N and C content was performed using an elemental analyzer (vario MACRO cube, Langenselbold, Germany).

2.5. ¹³C and ¹⁵N Analysis

The leaf ¹³C and ¹⁵N signatures were determined using an isotope ratio mass spectrometer (MAT253, Thermo Fisher Scientific, Langenselbold, Germany) coupled with an elemental analyzer. ¹³C/¹²C or ¹⁵N/¹⁴N values were calculated as follows relative to their international references:

$$\delta {[{}^{13}\mathrm{C},{}^{15}\mathrm{N}]}_{\mathrm{samples}}=(R_{\mathrm{sample}}/R_{\mathrm{standard}}-1)\times 1000$$

(3)

where R_sample is the ¹³C/¹²C or ¹⁵N/¹⁴N ratio, and R_standard is the international standard as VPDB for C (0.1‰) and atmospheric N₂ for N (0.2‰) [33].

2.6. Quantifying Δ¹⁵N Values of NH₄⁺ Assimilation and N Source Partitioning

For plantlets simultaneously assimilating both nitrate and ammonium, leaf δ¹⁵N values represents an integration of NO₃⁻- and NH₄⁺-derived N. A two end-member isotope mixing model [10,22] was employed to quantify their relative contributions:

$${\delta }_{\mathrm{T}}=f_{\mathrm{A}}{\delta }_{\mathrm{A}}+f_{\mathrm{B}}{\delta }_{\mathrm{B}}=f_{\mathrm{A}}{\delta }_{\mathrm{A}}+\left(1-f_{\mathrm{A}}\right){\delta }_{\mathrm{B}}$$

(4)

where δ_T represents the experimentally measured leaf δ¹⁵N of O. violaceus plantlets grown with mixed N sources; δ_A and δ_B denote the δ¹⁵N values of assimilates that arise from the NO₃⁻ and NH₄⁺ assimilation. The partitioning of NO₃⁻ is denoted f_A, and that of NH₄⁺ is denoted f_B. Since these are complementary, f_B = 1 − f_A.

While NH₄⁺ can induce toxicity when supplied as the sole N source, NO₃⁻ generally does not inhibit growth. To investigate this contrast, we applied two experimental treatments utilizing light (L) and heavy (H) stable N isotope-labeled NO₃⁻. This approach enabled the derivation of f_A and f_B, representing the fractional contributions of NH₄⁺- and NO₃⁻-derived N, respectively, to plant assimilation. The resulting calculations are as follows:

$${\delta }_{\mathrm{T}H}=f_{\mathrm{A}H}{\delta }_{\mathrm{A}H}+f_{\mathrm{B}}{\delta }_{\mathrm{B}}=f_{\mathrm{A}H}{\delta }_{\mathrm{A}H}+\left(1-f_{\mathrm{A}H}\right){\delta }_{\mathrm{B}}$$

(5)

$${\delta }_{\mathrm{T}L}=f_{\mathrm{A}L}{\delta }_{\mathrm{A}L}+f_{\mathrm{B}}{\delta }_{\mathrm{B}}=f_{\mathrm{A}L}{\delta }_{\mathrm{AL}}+\left(1-f_{\mathrm{A}L}\right){\delta }_{\mathrm{B}}$$

(6)

In this experiment, O. violaceus plantlets for both the light (L) and heavy (H) ¹⁵N isotope treatments were cultivated under uniform conditions with identical growth media. The sole experimental difference was the δ¹⁵N value of the supplied NO₃⁻ (denoted δ_L,NO3 and δ_H,NO3 for L and H treatments, respectively). Given that stable N isotopes are physiologically inert and do not influence plant metabolism, growth, or other biological parameters, the principle of isotopic equivalence was applied. This established that the fractional contribution of NO₃⁻-derived N to assimilation (f_A) remains constant across treatments (f_A = f_A,H = f_A,L). Consequently, the relationship is f_A = f_AH = f_AL, where 1 − f_AH = 1 − f_AL simplifies to the final equation:

$$f_{\mathrm{A}}=\left({\delta }_{\mathrm{T}H}-{\delta }_{\mathrm{T}L}\right)/\left({\delta }_{\mathrm{A}H}-{\delta }_{\mathrm{A}L}\right)$$

(7)

In this study, δ_TH denotes the leaf δ¹⁵N of O. violaceus plantlets grown with mixed N sources (NO₃⁻ and NH₄⁺), where the NO₃⁻ had a δ¹⁵N of 22.67‰ (high δ¹⁵N, H treatment). Conversely, δ_TL represents leaf δ¹⁵N value plantlets grown under identically mixed N sources, but with NO₃⁻ at lower δ¹⁵N = 8.08‰ (low δ¹⁵N, L treatment). The standard error (SE) of the fractional contribution parameter f_A was calculated using error propagation [34].

After determining f_A, δ_B (δ¹⁵N_B) can be calculated by Equation (5) or Equation (6) as follows:

$${\delta }_{\mathrm{B}}=\left({\delta }_{\mathrm{A}H}{\delta }_{\mathrm{T}L}-{\delta }_{\mathrm{A}L}{\delta }_{\mathrm{T}H}\right)/\left({\delta }_{\mathrm{A}H}+{\delta }_{\mathrm{T}L}-{\delta }_{\mathrm{A}L}-{\delta }_{\mathrm{T}H}\right)$$

(8)

The error propagation formula was also used to obtain the standard error (SE) of δ_B [34].

The N isotope discrimination of assimilates originating from leaf NH₄⁺ assimilation (Δ¹⁵N_B) relative to the source was calculated as follows [35]:

$${\Delta }^{15}{\mathrm{N}}_{\mathrm{B}}\left(\text{‰}\right)={\delta }^{15}{\mathrm{N}}_{\mathrm{ammonium}}-{\delta }^{15}{\mathrm{N}}_{\mathrm{B}}$$

(9)

where δ¹⁵N_ammonium was −2.64‰. The error propagation formula was further used to obtain the standard error (SE) of Δ¹⁵N_B [34].

In this study, both δ_TH and δ_TL were directly measured. However, determining the δ_AL and δ_AH parameters reflecting N isotope discrimination during NO₃⁻ assimilation and unassimilated NO₃⁻ translocation between shoots and growth medium, presented significant challenges under mixed N sources. Both δ_AL and δ_AH exhibited dynamically temporal changes throughout the experiment. Critically, reliable quantification of δ_AL and δ_AH was only achievable when plantlets were grown with NO₃⁻ as the sole N source.

While δ_AL and δ_AH in NO₃⁻-fed plantlets could be theoretically influenced by unassimilated NO₃⁻, prior studies indicate that this effect is negligible under the experimental conditions. For instance, prior studies indicate this effect is negligible under our experimental conditions. Studies in tomato and tobacco demonstrated that leaf NO₃⁻ storage pools are replenished during darkness and depleted in daylight, resulting in minimal leaf NO₃⁻ levels by afternoon [36,37]. Given that harvests occurred in the afternoon after five weeks of culture, unassimilated NO₃⁻ constituted a trivial fraction relative to assimilated NO₃⁻. Furthermore, leaf δ¹⁵N values remained consistent across 10 to 40 mM NO₃⁻ treatments [22,38], confirming that residual unassimilated NO₃⁻ exerted negligible effects on δ¹⁵N signatures. Consequently, δ_AL and δ_AH values for O. violaceus plantlets grown with mixed N sources were considered equivalent to those of NO₃⁻-fed plantlets.

In prior studies, where NaNO₃ served as the sole N source (δ¹⁵N = 8.08‰ or 22.67‰) [22,38], the mean leaf δ¹⁵N values of O. violaceus plantlets across 10, 20, and 40 mM NO₃⁻ concentrations closely aligned with δ¹⁵N values (δ_AL or δ_AH) in this study. Specifically, δ_AL (5.71 ± 0.17‰, n = 9) derived from low-δ¹⁵N (L) treatment [38], while δ_AH (17.02 ± 0.23‰, n = 9, SE) corresponded to high-δ¹⁵N (H) treatment [22]. Using these parameters (δ_TH, δ_TL, δ_AH and δ_AL), we calculated the proportional contributions of f_A and f_B of NO₃⁻- and NH₄⁺-derived N. Although NO₃⁻ efflux could theoretically alter δ_AL and δ_AH values by redistributing unassimilated NO₃⁻, leaf δ¹⁵N remained invariant across 10 mM to 40 mM NO₃⁻ concentrations [22,38]. This demonstrates that the presence of NH₄⁺ in mixed sources negligibly affects NO₃⁻ isotope discrimination, and efflux dynamics did not significantly perturb leaf isotopic signatures. Thus, δ_AL and δ_AH values from NO₃⁻-fed plantlets were valid proxies for mixed-N treatments. Nevertheless, minor uncertainties calculated via Equation (5) may persist due to unquantified efflux effects.

2.7. Quantifying the Absolute Nitrogen and Carbon Accumulation in Leaves

Leaf N accumulation amount (NAA) and C accumulation amount (CAA) were calculated using the following two equations:

$$\mathrm{N}\mathrm{A}\mathrm{A}=(\mathrm{D}\mathrm{W}\times \mathrm{N}\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{t})/\mathrm{M}$$

(10)

$$\mathrm{C}\mathrm{A}\mathrm{A}=(\mathrm{D}\mathrm{W}\times \mathrm{C}\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{t})/\mathrm{M}$$

(11)

where the molar mass of N or C is denoted as M.

2.8. Quantifying Leaf Nitrogen from NO₃⁻/NH₄⁺ Assimilation

The leaf NAA from NO₃⁻/NH₄⁺ assimilation are calculated using the following equations:

$${\mathrm{NAA}}_{\mathrm{nitrate}}=\mathrm{NAA}\times f_{\mathrm{A}}$$

(12)

$${\mathrm{NAA}}_{\mathrm{ammonium}}=\mathrm{NAA}\times f_{\mathrm{B}}$$

(13)

where the error propagation formula was used to calculate the standard error (SE) of NAA_nitrate or NAA_ammonium [34].

2.9. Statistical Analysis

Data are presented as means ± standard errors (SE n = 3). Statistical significances were determined using analysis of variance (ANOVA), with treatment means compared by Tukey’s honestly significant difference (HSD) test (p < 0.05). All analyses were performed using a Data Processing System (DPS) software (version 7.05, Hangzhou Ruifeng Information Technology Co., Ltd., Hangzhou, China).

3. Results

3.1. Growth Characteristics

The growth of O. violaceus plantlets was significantly influenced by NO₃⁻ concentrations under a constant 20 mM NH₄⁺. Increasing NO₃⁻ concentrations substantially enhanced plantlet growth. Biomass and leaf dry weight declined significantly only at the lowest tested NO₃⁻ concentration (

Table 1. Effects of nitrate levels on biomass production in 35-day-old Orychophragmus violaceus plantlets grown under a combined inorganic nitrogen source with a fixed 20 mM NH₄-N.

Parameters	Inorganic Nitrogen Concentration
	10 mM NO3-N + 20 mM NH4-N	20 mM NO3-N + 20 mM NH4-N	40 mM NO3-N + 20 mM NH4-N
Increased biomass (g/plantlet FW)	2.525 ± 0.084 b	4.178 ± 0.369 a	4.070 ± 0.251 a
Leaf DW (g)	0.060 ± 0.003 b	0.107 ± 0.006 a	0.111 ± 0.008 a

Data indicate means ± SE (n = 3). Values in the same row followed by different letters are significantly different at p < 0.05.

), indicating that insufficient NO₃⁻ availability severely limited growth under these conditions.

3.2. Chlorophyll Concentrations

Chlorophyll levels in O. violaceus plantlets were significantly influenced by NO₃⁻ concentration. Under a constant 20 mM NH₄⁺, increasing NO₃⁻ substantially supplied elevated chlorophyll concentrations. Levels increased over threefold as NO₃⁻ concentrations rose from 10 to 20 mM (

Table 2. Effects of nitrate levels on chlorophyll concentration in 35-day-old Orychophragmus violaceus plantlets grown under a combined inorganic nitrogen source with a fixed 20 mM NH₄-N.

Parameters	Inorganic Nitrogen Concentration
	10 mM NO3-N + 20 mM NH4-N	20 mM NO3-N + 20 mM NH4-N	40 mM NO3-N + 20 mM NH4-N
Chl a (mg/g FW)	0.163 ± 0.008 b	0.564 ± 0.037 a	0.647 ± 0.047 a
Chl b (mg/g FW)	0.081 ± 0.005 b	0.295 ± 0.013 a	0.322 ± 0.012 a
Chl a + b (mg/g FW)	0.243 ± 0.004 b	0.859 ± 0.049 a	0.969 ± 0.057 a

Data indicate means ± SE (n = 3). Values in the same row followed by different letters are significantly different at p < 0.05.

). This enhancement in chlorophyll biosynthesis was strongly associated with elevated NO₃⁻ availability, highlighting its critical role in supporting photosynthetic capacity.

3.3. Leaf Nitrogen and Carbon Content

While increasing NO₃⁻ concentrations had no significant effects on leaf N content in O. violaceus plantlets, which remained above 6% across all treatments, they significantly influenced leaf C content. Elevated NO₃⁻ supply significantly increased leaf C, with the lowest concentration consistently yielding reduced C content compared to higher concentrations (Figure 1).

3.4. Leaf Carbon Isotope Composition

While NO₃⁻ concentration significantly influenced δ¹³C values of O. violaceus plantlets, significant differences occurred only at the lowest NO₃⁻ concentration. δ¹³C values remained stable across the 20 to 40 mM NO₃⁻ range (Figure 2).

3.5. Leaf Nitrogen Isotope Composition

δ¹⁵N values in O. violaceus plantlets were significantly higher in the H treatment than in the L treatment (Figure 3). Increasing NO₃⁻ concentration significantly elevated δ¹⁵N values in both the L and H treatments. Notably, under the lowest 10 mM NO₃⁻ with mixed N source, δ¹⁵N values in the L treatment were below −3.0‰. This ¹⁵N depletion relative to the N source (−2.64‰) suggests isotopic discrimination during NH₄⁺ assimilation.

3.6. The Proportion of Inorganic Nitrogen Utilization

Nitrate concentration had no significant effect on NO₃⁻ utilization in O. violaceus plantlets (Figure 4). Increasing NO₃⁻ levels failed to increase NO₃⁻ uptake, as NH₄⁺ remained the preferred N source, evidenced by its significantly higher utilization proportion.

3.7. The δ¹⁵N and Δ¹⁵N of Ammonium Assimilation

The δ¹⁵N values of NH₄⁺ assimilation in the O. violaceus plantlets were consistently depleted relative to the source δ¹⁵N across all treatments (Figure 5a), indicating isotopic discrimination during assimilation. However, the magnitude of this discrimination (Δ¹⁵N) varied significantly with NO₃⁻ concentration. Increasing NO₃⁻ levels significantly reduced Δ¹⁵N values (Figure 5b), demonstrating an inverse relationship between NO₃⁻ availability and isotopic discrimination during NH₄⁺ assimilation.

3.8. Absolute Nitrogen and Carbon Accumulation in the Leaves

Leaf N accumulation amount (NAA) in O. violaceus plantlets increased with NO₃⁻ availability. Higher NO₃⁻ levels significantly enhanced both leaf NAA (Figure 6a) and C accumulation amount (CAA) (Figure 6b). Critically, plantlets grown at the lowest NO₃⁻ concentration exhibited significantly lower NAA and CAA compared to higher concentrations.

3.9. Leaf Nitrogen Accumulation Amount Resulting from Nitrate/Ammonium Assimilation

Elevated NO₃⁻ concentrations enhanced both NO₃⁻ and NH₄⁺ assimilation in O. violaceus plantlets. Leaf NAA from NO₃⁻ (NAA_nitrate) and NH₄⁺ (NAA_ammonium) increased progressively with higher NO₃⁻ concentrations (Figure 7). Strikingly, doubling NO₃⁻ from 10 to 20 mM significantly boosted both NAA_nitrate and NAA_ammonium, demonstrating its role as a key driver of N assimilation efficiency.

4. Discussion

Ammonium is generally preferred over NO₃⁻ as a N source due to its lower energy cost during assimilation [1]. However, elevated NH₄⁺ concentrations typically induce growth suppression (NH₄⁺ toxicity) in plants [4,39,40]. While supplying mixed NO₃⁻ and NH₄⁺ can alleviate this toxicity [5,8], the precise mechanisms remain unresolved. Notably, NH₄⁺ toxicity symptoms often persist even in the presence of NO₃⁻ [10,41,42]. In this study, O. violaceus plantlets supplemented with 20 mM NH₄⁺ exhibited a significant decrease in biomass and chlorophyll concentrations at 10 mM NO₃⁻ compared to higher NO₃⁻ treatments (Table 1 and Table 2), despite maintaining leaf N levels > 6% across all treatments. These results contrast with the well-documented positive correlations between chlorophyll and leaf N content, where ~75% of leaf N is allocated to chloroplasts [43]. These findings suggest that the capacity of NO₃⁻ to mitigate NH₄⁺ toxicity depends on its concentration and may involve mechanisms independent of total N status.

Excessive NH₄⁺ concentrations in growth solutions can induce futile NH₄⁺ cycling [12,44], a process characterized by disproportionately high NH₄⁺ efflux relative to influx. This efflux energetically costly [11,45], leading to significant energy drain that impedes biomass accumulation—a hallmark of NH₄⁺ toxicity [46,47]. Crucially, because effluxed NH₄⁺ is enriched in ¹⁵N [31], the futile NH₄⁺ cycling results in ¹⁵N-depleted assimilates during NH₄⁺ metabolism. The magnitude of this depletion, quantified as Δ¹⁵N_B, serves as a proxy for futile cycling activity.

In O. violaceus plantlets grown with mixed N sources, δ¹⁵N_B values of NH₄⁺ assimilation were consistently more negative than −2.64‰ (Figure 5a), confirming NH₄⁺ efflux across all treatments. Notably, Δ¹⁵N_B values decreased with increasing NO₃⁻ concentrations (Figure 5b), suggesting that elevated NO₃⁻ availability suppresses futile NH₄⁺ cycling. These results imply that NO₃⁻-mediated mitigation of NH₄⁺ toxicity operates, at least partly, through suppression of futile cycling, with efficacy critically dependent on NO₃⁻ concentration.

To mitigate the detrimental effects of high NH₄⁺ concentrations, plants often expel excess NH₄⁺ or enhance its assimilation [26,27]. In O. violaceus plantlets, leaf N derived from NH₄⁺ assimilation (NAA_ammonium) increased significantly with elevated NO₃⁻ concentrations (Figure 7), indicating that higher NO₃⁻ availability stimulated NH₄⁺ assimilation [48]. Given the constant 20 mM NH₄⁺ supply across treatments, this rise in NAA_ammonium reflected improved NH₄⁺ utilization efficiency. This enhanced efficiency likely suppressed futile NH₄⁺ cycling—a key driver of NH₄⁺ toxicity [47].

However, excessive NH₄⁺ assimilation can result in intracellular and extracellular acidification, a primary driver of NH₄⁺ toxicity [13]. Notably, in this study, increased NH₄⁺ assimilation coincided with enhanced NO₃⁻ assimilation (Figure 7). Crucially, NO₃⁻ reduction consumes protons [1,13], generating hydroxyl ions that neutralize acidity. Concurrently, NO₃⁻ uptake via NO₃⁻/H⁺ symporters [21,49,50] further alleviates acidic stress by co-transporting protons into cells. Thus, NO₃⁻ assimilation not only improves N utilization but also directly counterbalances acidification from NH₄⁺ assimilation. These findings reveal a dual mechanism for NO₃⁻-mediated mitigation of NH₄⁺ toxicity: (1) suppression of futile NH₄⁺ cycling through enhanced assimilation efficiency, and (2) neutralization of acidic stress via proton consumption during NO₃⁻ reduction and proton-coupled transport.

While NO₃⁻ reduction can alleviate acidic stress, insufficient NO₃⁻ availability may fail to effectively mitigate this response [51]. In O. violaceus plantlets, the chlorophyll concentrations significantly decreased at 10 mM NO₃⁻ compared to higher NO₃⁻ concentrations (Table 2), despite leaf N content reaching its maximum at this same 10 mM NO₃⁻. This paradoxical reduction in chlorophyll under lower NO₃⁻ likely stems from acidic stress induced by excessive NH₄⁺ assimilation [13].

Chlorophyll concentration serves as an indirect indicator of photosynthetic capacity, as reduced levels directly impede plant photosynthetic potential [52]. In this study, O. violaceus plantlets grown at 10 mM NO₃⁻ exhibited the lowest chlorophyll content, indicating severely compromised photosynthetic capacity. These O. violaceus plantlets were grown mixotrophically, utilizing both CO₂ and sucrose.

Carbon isotope discrimination during key metabolic processes was characterized as follows. (1) Photosynthetic CO₂ assimilation: Discrimination averaged ~20‰ [25]. Given the δ¹³C value of chamber CO₂ was −10.55 ± 0.13‰ (n = 12), the δ¹³C value of assimilated CO₂-derived C is estimated −10.55 + (−20 ‰) = −30.55‰. (2) Sucrose utilization, Discrimination (fractionation) was 2.54 ± 0.13‰ (n = 3) [10]. Given the δ¹³C value of supplied sucrose was −11.64‰, the δ¹³C value of metabolized sucrose-derived C is calculated −11.64 + (−2.54‰) = −14.18‰.

Leaf δ¹³C values reflect the integrated δ¹³C signatures of assimilated CO₂ and metabolized sucrose. Greater reliance on inorganic CO₂ assimilation results in lower (more negative) leaf δ¹³C values due to photosynthetic fractionation, while increased utilization of organic sucrose elevates leaf δ¹³C [24]. As shown in Figure 2, O. violaceus plantlets grown at 10 mM NO₃⁻ exhibited significantly higher leaf δ¹³C values than those at other NO₃⁻ concentrations. These indicate a substantial reduction in photosynthetic CO₂ assimilation relative to sucrose utilization. These isotopic data align with the observed photosynthetic deficiency and corroborate the impaired photosynthetic capacity at this NO₃⁻. We conclude that insufficient chlorophyll synthesis at 10 mM NO₃⁻ directly constrains photosynthetic performance through reduced CO₂ assimilation.

As previously established, mitigating NH₄⁺ toxicity in plants—whether through extruding excess NH₄⁺ or enhancing its assimilation—demands substantial energy [4,26,27,53,54]. In this study, the O. violaceus plantlets grown at the lowest 10 mM NO₃⁻ concentration exhibited the highest leaf N content (>6.5%, Figure 1a), with ~70% of assimilated N derived from NH₄⁺ (Figure 4b). However, this high-N phenotype incurred significant energy costs, exacerbated by pronounced futile NH₄⁺ cycling—an ATP-intensive process [11]. Concurrently, leaf C content decreased markedly at 10 mM NO₃⁻ (Figure 1b), reflecting a metabolic trade-off likely driven by energy depletion [55].

Despite exogenous sucrose supplementation (3% w/v), energy limitation persisted in low-NO₃⁻ conditions due to impaired photosynthetic capacity [44]. Increased respiratory activity from sucrose catabolism may further exacerbate oxidative stress via mitochondrial reactive oxygen species generation [8,56], explaining the stunted growth at 10 mM NO₃⁻ (Table 1). Thus, even 10 mM NO₃⁻ in mixed N sources failed to fully mitigate NH₄⁺ toxicity, as sucrose-derived energy proved insufficient to meet metabolic demands of NH₄⁺ detoxification.

Notably, photosynthetic capacity significantly improved as NO₃⁻ increased from 10 to 20 mM (Figure 2). Enhanced photosynthesis likely bolsters cellular energy production through chloroplast electron transport—where the noncyclic pathways generate both ATP and NADPH, while the cyclic pathways produce ATP exclusively [57]. This dynamic modulation of ATP:NADPH output better aligns with the stoichiometric demands of NH₄⁺ detoxification, enabling efficient assimilation without depleting C reserves.

5. Conclusions

Our study demonstrates that futile NH₄⁺ cycling occurs in plants under NH₄⁺ toxicity stress, with its magnitude modulated by NO₃⁻ availability. Elevated NO₃⁻ concentrations enhance NH₄⁺ utilization efficiency, suppressing futile cycling and its associated energy drain. Concurrently, increased NO₃⁻ assimilation alleviates acidic stress via proton-consuming reduction and transport, counteracting acidification from NH₄⁺ assimilation. Crucially, the lowest NO₃⁻ concentration (10 mM) proved insufficient to mitigate acidic stress, which results in an inhibition of chlorophyll biosynthesis. The chlorophyll deficiency caused by acidic stress severely impaired photosynthetic capacity, which serves as a core mechanism of NH₄⁺ toxicity. Thus, NO₃⁻-mediated mitigation of NH₄⁺ toxicity operates through dual pathways: (1) restoring photosynthetic energy production and (2) enhancing NH₄⁺ utilization efficiency to minimize futile cycling.