Growth conditions and experimental design

Seeds of Brassica napus cv. ES Neptune (Euralis Semillas S.A.) were sterilized by immersion in 70% ethanol for 30 s, then in 1% bleach for 7 min and finally rinsed 5 times with deionised water. Seeds were sown in trays filled with perlite-vermiculite 1:1 (v/v) inert substrate mixture and watered with deionised water. Trays were kept during 4 days in the dark at 4 °C for vernalization and transferred into a phytotron [60/70% of humidity, 23 °C day (14 h) with a light intensity of 400 μmol m⁻² s⁻¹ and 18 °C night (10 h)].

Seven day-old seedlings were washed with deionised water to remove traces of perlite-vermiculite and transferred to 5 L hydroponic tanks (10 plants per tank; for Additional file 1: Fig. S1 20 plants per tank). The nutrient solution contained 1.15 mM K₂HPO₄, 0.85 mM MgSO₄, 0.7 mM CaSO₄, 2.68 mM KCl, 0.5 mM CaCO₃, 0.07 mM NaFeEDTA, 16.5 μM Na₂MoO₄, 3.7 μM FeCl₃, 3.5 μM ZnSO₄, 16.2 μM H₃BO₃, 0.47 μM MnSO₄, 0.12 μM CuSO₄, 0.21 μM AlCl₃, 0.126 μM NiCl₂ and 0.06 μM KI, pH 6.8. Two concentrations of N were tested for each source: 0.5 and 1 mM. To properly compare both N sources within each concentration studied, NO₃ ⁻-fed plants were supplied with CaSO₄ to equilibrate the SO₄ ⁻² supplied together with the NH₄ ⁺. Thus, for 0.5 mM N treatment we applied 0.25 mM SO₄(NH₄)₂ for ammonium-fed plants and 0.25 mM Ca(NO₃)₂ + 0.25 mM CaSO₄ for nitrate-fed ones. Similarly, for 1 mM N treatment we applied 0.5 mM SO₄(NH₄)₂ for ammonium-fed plants and 0.5 mM Ca(NO₃)₂ + 0.5 CaSO₄ for nitrate-fed ones. It should be noted that plants grown with 0.5 mM and 1 mM N were supplied with 1.804 mM and 2.054 mM of total sulfur, respectively. Similarly, for Additional file 1: Figure S1, the sulfate supplied in the nutrient solution was 1.804, 2.805 and 4.055 mM for 0.5, 2.5 and 5 mM N treatments, respectively. Plants were maintained in hydroponic culture for 18 days. The nutrient solution of the tanks was changed on days 7, 10 and 14. The pH of the solution was monitored every two days and maintained stable between 6.8 and 7.2 during the whole experiment. A total of four tanks were used per treatment, with ten plants per tank. Thus, a total of 40 plants were grown per condition.

Harvest

Chlorophyll content and gas exchange parameters in two plants per tank were measured in the first fully expanded leaf using a SPAD-502 Meter and an open-flow portable photosynthesis system (LICOR 6400), respectively. Chlorophyll was determined in every plant and gas exchange parameters in two plants per tank. For gas exchange, a 6 cm² leaf area chamber was used making the measurements at 60% of humidity and 400 ppm CO₂. After in vivo measurements, shoots and roots were separated and individually weighed. Then, all the plants grown in the same tank were pooled, frozen in liquid nitrogen and homogenized in a Tissue Lyser (Retsch MM 400). Samples were stored at −80 °C until use.

Element and metabolite determination

Nitrogen, carbon and sulfur content were determined by combustion of plant dry material using an elemental analyser Flash EA1112 (Thermo Fisher Scientific Inc., Waltham, MA, USA).

Individual amino acids, ammonium, sulfate, Cys and GSH were extracted from 100 mg of frozen leaf and root powder as described in Sarasketa et al. (2014) [28]. Ammonium was quantified following the phenol hypochlorite method. Sulfate was quantified by capillary electrophoresis (Agilent G1600 CE3D, Agilent Technologies, Santa Clara, CA, USA) with indirect UV detection at 350 nm and 240 nm reference according to the standard EPA Test Method 6500 (www.epa.gov). The capillary was 40 cm long with 10 μm internal diameter and the buffer used was the Agilent Life Sciences Inorganic anion buffer pH 7.7 (Ref. 8500–6797). Nitrate content was determined spectrophotometrically from aqueous extracts according to the method reported by Cataldo et al. (1975) [29].

For amino acid and GSH determination, extracts were neutralized with NaOH and after derivatization with 1 mM fluorescein isothiocyanate, amino acids were quantified by capillary electrophoresis (PA-800, Beckman Coulter Inc., USA) coupled to laser-induced fluorescence detection (argon laser at 488 nm) as previously described [30]. Cysteine and GSH content were determined from the same extracts derivatized with 5-iodoacetamide fluorescein and reduced with tributylphosphine [31].

Glucosinolates were extracted by adding 1 mL of MeOH:water (70:30) to 25 or 50 mg of frozen leaf or root powder, respectively. The mixtures were homogenized with 2 mm glass beads in a Tissue Lyser (Retsch MM 400), incubated for 15 min at 80 °C to inactivate myrosinase and then, centrifuged for 20 min at 14000 g. Glucosinolates were determined from supernatants by UHPLC-QTOFMS analyses using an Acquity UPLC^TM from Waters (Milford, MA) interfaced to a Synapt G2 QTOF from Waters with electrospray ionization as described in Glauser et al. (2012) [32]. Glucosinolates were quantified using glucoraphanin and glucobrassicin as standards.

Protein extraction and enzyme activities

For protein quantification total soluble protein was extracted from 100 mg of frozen leaf or root powder with 1 mL of extraction buffer as described in Sarasketa et al. (2016) [6]. Protein was quantified using a Bradford-base dye-binding assay (Bio-Rad, Hercules, CA, USA) with bovine serum albumin as a standard. Every enzyme was determined with a 96-well plate spectrophotometer (BioTek Instruments). Glutamine synthetase (GS), NADH-glutamine 2-oxoglutarate aminotransferase (NADH-GOGAT), TCA anaplerotic enzymes and glutamate dehydrogenase (GDH) in its aminating sense were determined as described in Sarasketa et al. (2016) [6]. ATP sulfurylase (ATPS) activity was measured according to Lappartient and Touraine (1996) [33]. The pyrophosphate formed during the reaction was determined at 820 nm as described by Katewa and Katyare (2003) [34]. For O-acetylserine (thiol) lyase (OASTL) activity, the method described by Pajuelo et al. (2007) [35] was followed quantifying the Cys formed during the reaction with ninhydrin reagent at 560 nm as described by Gaitonde (1967) [36].

RNA extraction and qPCR

RNA was extracted from 50 mg of leaves or roots with the Nucleospin RNA plant kit (Macherey-Nagel), which includes DNAse treatment, and 1 μg of RNA was retrotranscribed into cDNA (PrimeScriptTM RT; Takara Bio Inc.). The absence of contamination with genomic DNA was tested by quantitative RT-PCR in all RNA samples. Gene expression was determined from 2 μL of cDNA diluted 1:10 in a 15 μL reaction volume using SYBR Premix ExTaqTM (Takara Bio Inc.) in a Step One Plus Real Time PCR System (Applied Biosystems). The primers used for amplifying SULTR1;1 (AJ416460) and SULTR1;2 (AJ311388) were described in Abdallah et al. (2010) [37]. For amplifying SULTR1;3 (XM_013896883), the primers used were forward 5′-TTGATTGATTTCTTATCCCATGC-3′ and reverse 5′-GGAACCCTTTGAGTTGTTGAA-3′; for SULTR2;1 (NM_001315588) forward 5′-TCTTGCAAAGCTTGATCCTC -3′ and reverse 5′-CGATTGCTATCTCTCTTGATG-3′ and for SULTR2;2 (XM_013840590) forward 5′-TTTCAAGCCATCTTTGGACTC-3′ and reverse 5′-AGCCATGAACCCAACAAGAG-3′. The PCR program was: 95 °C for 5 min, 40 cycles of 15 s at 94 °C followed by 1 min at 60 °C; finally, a melting curve was programmed (40–95 °C with one fluorescence read every 0.3 °C). Relative gene expression was calculated as the ΔCp between each gene and the average of the housekeeping genes ACTIN (AF111812) with the primers forward 5′-GATTCCGTTGCCCTGAAGTA-3′ and reverse 5′-GCGACCACCTTGATCTTCAT-3′ and EF1-a (DQ312264) with the primers forward 5′- GCCTGGTATGGTTGTGACCT-3′ and reverse 5′ GAAGTTAGCAGCACCCTTGG-3′. The stability of the reference genes across samples was tested using geNorm software [38]. The efficiency for every primer couple was calculated with serial cDNA dilutions. The efficiencies were 2.1 for SULTR1;1, 1.9 for SULTR1;2, 2.1 for SULTR1;3, 2.0 for SULTR2;1 and 2.0 for SULTR2;2. Data are presented respect to the highest value obtained within each tissue (value “1”).

Statistical analysis

All the data presented are given as means with standard errors. Data were analyzed using SPSS 17.0 (Chicago, IL, USA). Statistical analysis of normality and homogeneity of variance were analyzed by Kolmogorov-Smirnov and Levene tests. The significance of the results was assessed using independent samples t-test and by ANOVA analysis followed by Duncan’s post hoc test.

Nitrogen assimilation is enhanced in leaves of ammonium-fed B. napus plants

A large number of studies have associated both N availability and plant N status with the regulation of S uptake and assimilation [1, 2]. However, the studies taking into account N source effect are scarce. In this work, we studied N and S metabolism in rapeseed plants hydroponically grown with NO₃ ⁻ or NH₄ ⁺ as sole N source. Ammonium nutrition may be a stressful situation and its effects range from an induction in NH₄ ⁺ assimilatory machinery to even plant death in function of the severity of the stress. Chlorosis is considered as a symptom of severe ammonium stress while chlorophyll accumulation has been reported as a plant response upon mild ammonium stress [43, 44]. In this work, we aimed to study N source effect avoiding severe ammonium toxicity and therefore we chose 0.5 and 1 mM N concentrations, conditions where photosynthesis or chlorophyll content remained unaffected (Table 1). Still, it was evident observing plant growth, a higher severity of the stress in A1 compared to A0.5 (Fig. 1). This situation was confirmed with NH₄ ⁺ content, considered as a classical metabolic marker of ammonium stress in tissues, as only in A1 leaves increased compared to N1 leaves (Fig. 2b) while no differences were observed at 0.5 mM. Taken together, we can assume that oilseed rape plants were exposed to mild and moderate ammonium stress conditions in A0.5 an A1, respectively.

Scavenging excessive NH₄ ⁺ into N-containing molecules is considered as a strategy to prevent ammonium toxicity [4, 5] and this strategy seems essential to prevent unrestrained NH₄ ⁺ accumulation in B. napus (Fig. 2b). Indeed, NH₄ ⁺ assimilation was clearly enhanced in ammonium-fed plants and notably in A1 leaves, which was reflected by higher total N, free amino acid and protein content compared to N1 leaves (Fig. 2a–d). In agreement with the higher content of N-compounds observed in ammonium-fed plants, GS and NADH-GOGAT activities increased under ammonium nutrition (Fig. 3a, b). The induction of GS/GOGAT has been commonly reported in plants grown with NH₄ ⁺ as N source, including Arabidopsis thaliana [6, 45]. Moreover, the use of Arabidopsis mutants has revealed the importance of these enzymes in NH₄ ⁺scavenging [5, 45]. NAD(H)-GDH activity or gene expression induction is also a classical response of plants exposed to ammonium stress including species of the Brassicaceae family [6, 46] (Fig. 3c). NADP(H)-GDH enzyme has been poorly characterized in plants. It is interesting that this enzyme is also induced in leaves of ammonium-fed rapeseed plants (Fig. 3d). Indeed, the induction of both NAD(H)- and NADP(H)-dependent GDH was also reported in other situations where internal NH₄ ⁺ accumulated, such as in Lotus japonicus plants deficient in plastidic GS under active photorespiratory conditions [47]. Although studied for long time, the role of GDH in plant metabolism remains unresolved; some works indicate an exclusive GDH role for 2-OG provision [48] while in other reports an aminating role is suggested [49]. Under ammonium stress, it is somehow logic to envision an aminating function that could contribute to avoid NH₄ ⁺ rise to toxic levels. However, 2-OG provision for amino acid synthesis is also crucial and thus, GDH could be working deaminating Glu. It will be interesting to study in the future whether NAD(H)- and NADP(H)-dependent GDH enzymes have a differential function in plant metabolism in general and under ammonium stress in particular. In line with the role of GDH to maintain organic acids pool, anaplerotic Tricarboxylic acid cycle (TCA) enzymes have also been suggested as important to favor NH₄ ⁺ assimilation [6, 50]. In the present work, an induction of TCA enzymes was also observed in B. napus plants grown under ammonium nutrition (Additional file 3: Figure S3). Overall, it seems that the growth impairment observed under ammonium nutrition (Fig. 1) is not due to NH₄ ⁺ accumulation in tissues but rather to the energy cost associated to maintaining NH₄ ⁺ homeostasis by increasing its assimilation.

Sulfur metabolism is induced in leaves of B. napus plants grown under ammonium nutrition

Sulfur uptake and assimilation are strongly interconnected with N and C metabolism [51]. In Brassicaceae, glutathione (GSH) and glucosinolates (GLS) are the major S-containing biomolecules. In leaves of NH₄ ⁺-fed rapeseed plants the increase in both total GLS and total glutathione pool (GSH + GSSG) indicates that sulfur metabolism was enhanced (Fig. 4c, d). Both types of molecules are derived from amino acids, GSH from Cys, Glu and Gly and GLS mainly from Trp, Met and Phe. Indeed, we observed a general increase in free amino acid content (Fig. 2c), including Glu, Gly, Trp and Phe, in A1 leaves, together with Ser (Table 2), precursor of Cys and Met; thus, suggesting an increase in the flux of these amino acids towards GSH and GLS synthesis. According to this, ATPS activity, which catalyzes the first step of primary SO₄ ⁻² assimilation, and OASTL activity, which catalyzes the synthesis of Cys and, therefore, is the starting point for the synthesis of organic S-compounds, were induced in leaves of A1 plants (Fig. 5). Several works have also reported that the increase in amino acid availability enhanced SO₄ ²⁻ assimilation, for example through an increase of the incorporation of ³⁵S into proteins in Lemna minor [12] and in roots of A. thaliana [1]. Similarly, Zhu et al. (2006) [52] reported the increased expression of genes encoding S-assimilating enzymes together with a higher content of Cys and GSH in leaves of ammonium-fed rice compared to nitrate-fed one. Interestingly, in oilseed rape leaves Ser was, together with Glu, the most abundant amino acid (Table 2) and Zhu et al. (2006) [52] hypothesized that Ser, substrate of OAS to form Cys, might be the factor stimulating S assimilation under ammonium nutrition. In general, the accumulation of N in leaves under ammonium nutrition led to an increase in the availability of amino acids that was probably behind the stimulation of S incorporation into biomolecules. On the other hand, the activation of GLS and GSH synthesis could also be part of the plant strategy to store N when cell NH₄ ⁺ availability is excessive. These results obtained in oilseed rape are in line with those reported in a recent study from our research group where we showed an increase in the synthesis of aliphatic and indolic GLS in shoots of A. thaliana and broccoli under mild ammonium stress [27]. Similarly, Zaghdoud et al. (2016) [26] also observed higher levels of indolic GLS in broccoli plants under high ammonium stress associated to Trp accumulation. Another study focused on NO₃ ⁻/NH₄ ⁺ ratio effect on Chinese kale, showed that the presence of NH₄ ⁺ favored GLS synthesis [25]. In contrast, other works reported no changes or even decrease in GLS content when cultured with NH₄ ⁺ as sole N source [23, 24]. Therefore, it seems that GLS content is dependent on the culture conditions, the Brassica species/genotype and the degree of ammonium stress.

The increase in GLS level has also been observed in response to other environmental factors such as salinity [53] and water deficit [54]. For instance, Martínez-Ballesta et al. (2015) [55] reported a higher impact of salt stress in Arabidopsis mutants which had lost the capacity to synthesize aliphatic GLS. Similarly, the perturbation of aliphatic GLS synthesis by RNA interference (RNAi) caused a deregulation in different metabolites and proteins related to oxidative stress and photosynthesis [56]. Therefore, we could speculate that the increase in GLS level under ammonium nutrition, a part of being a consequence of the high availability of N-reduced compounds, could be a plant response upon abiotic stress. However, the molecular mechanism by which GLS metabolism is involved in the response of Brassicaceae family to abiotic stresses still needs to be deciphered.

Interestingly, in A1 leaves the stimulation of S assimilation was accompanied by SO₄ ²⁻ content diminution. However, this diminution cannot be fully explained by its recruitment into organic compounds. Sulfur is mainly taken up from the soil by SULTR SO₄ ²⁻ transporters. Arabidopsis encodes 12 SULTRs which are distributed in four groups (SULTR1, SULTR2, SULTR3 and SULTR4) [57]. In this work, we examined the expression of genes from SULTR1 and SULTR2 groups, located in the plasma membrane, to try to understand the variation in SO₄ ²⁻ content observed in function of the N source mainly at 1 mM concentration (Fig. 6). In roots SULTR1;2, the SULTR1 member with the highest expression, was induced in ammonium-fed plants (Fig. 6a), which is in agreement with the higher SO₄ ²⁻ content observed in this organ (Fig. 4b). Interestingly, in A1 leaves the down-regulation of the three types of SULTR1 transporters (Fig. 5b) is in line with the lesser SO₄ ²⁻accumulation in this organ (Fig. 4b). However, the accumulation of S-compounds in leaves evidences that SO₄ ²⁻ availability would not be limiting probably due to the high SO₄ ²⁻ content characteristic of cruciferous plants. The gene expression of SULTR1 transporters is commonly induced upon S deprivation [37, 40]; however, this induction is circumvented with the external supply of reduced S [40]. Indeed, SO₄ ²⁻ uptake is commonly repressed when reduced S is available, as for instance with the exogenous supply of GSH [33, 58] or H₂S [59]. Moreover, the inhibition of SO₄ ²⁻ uptake upon H₂S supply has even been observed in S-deficient plants [59]. Thus, in our study, the lower expression of SULTR1 transporters in A1 leaves seems to be related to the accumulation of sulfur-containing compounds rather than to the lower SO₄ ²⁻ accumulation. The overall data suggest a direct control of the N source on SO₄ ²⁻ uptake/transport by SULTR1 in leaves or most probably an indirect control by a negative feedback loop triggered by the accumulation of reduced S-compounds in line with the hypothesis of demand-driven regulation of S metabolism [33, 51].