Journal of Plant Physiology 171 (2014) 549–558

Contents lists available at ScienceDirect

Journal of Plant Physiology journal homepage: www.elsevier.com/locate/jplph

Physiology

Long-term sulphur starvation of Arabidopsis thaliana modifies mitochondrial ultrastructure and activity and changes tissue energy and redox status Monika Ostaszewska, Izabela M. Juszczuk ∗ , Izabella Kołodziejek, Anna M. Rychter Institute of Experimental Plant Biology and Biotechnology, Faculty of Biology, University of Warsaw, Miecznikowa 1, 02-096 Warsaw, Poland

a r t i c l e

i n f o

Article history: Received 3 September 2013 Received in revised form 12 December 2013 Accepted 16 December 2013 Keywords: Energy status Redox homeostasis Respiration Sulphur deficiency Mitochondria ultrastructure

s u m m a r y Sulphur, as a constituent of amino acids (cysteine and methionine), iron–sulphur clusters, proteins, membrane sulpholipids, glutathione, glucosinolates, coenzymes, and auxin precursors, is essential for plant growth and development. Absence or low sulphur concentration in the soil results in severe growth retardation. Arabidopsis thaliana plants grown hydroponically for nine weeks on Knop nutrient medium without sulphur showed morphological symptoms of sulphur deficiency. The purpose of our study was to investigate changes that mitochondria undergo and the role of the highly branched respiratory chain in survival during sulphur deficiency stress. Ultrastructure analysis of leaf mesophyll cells of sulphurdeficient Arabidopsis showed heterogeneity of mitochondria; some of them were not altered, but the majority had swollen morphology. Dilated mitochondria displayed a lower matrix density and fewer cristae compared to control mitochondria. Disintegration of the inner and outer membranes of some mitochondria from the leaves of sulphur-deficient plants was observed. On the contrary, chloroplast ultrastructure was not affected. Sulphur deficiency changed the respiratory activity of tissues and isolated mitochondria; Complex I and IV capacities and phosphorylation rates were lower, but external NAD(P)H dehydrogenase activity increased. Higher external NAD(P)H dehydrogenase activity corresponded to increased cell redox level with doubled NADH/NAD ratio in the leaf and root tissues. Sulphur deficiency modified energy status in the tissues of Arabidopsis plants. The total concentration of adenylates (expressed as ATP + ADP), measured in the light, was lower in the leaves and roots of sulphur-deficient plants than in the controls, which was mainly due to the severely decreased ATP levels. We show that the changes in mitochondrial ultrastructure are compensated by the modifications in respiratory chain activity. Although mitochondria of Arabidopsis tissues are affected by sulphur deficiency, their metabolic and structural features, which readily reach new homeostasis, make these organelles crucial for adaptation of plants to survive sulphur deficiency. © 2013 Elsevier GmbH. All rights reserved.

Introduction Sulphur (S) deficiency in soil is exerting deleterious effects on agriculture worldwide (Lewandowska and Sirko, 2008). Sulphate (SO4 2− ) uptake from the soil and incorporation into cysteine, a S-containing amino acid, are high energy-consuming processes. SO4 2− transporters in the root cells hydrolyse ATP to move

Abbreviations: AOX, alternative oxidase; BSA, bovine serum albumin; DW, dry weight; EDTA, ethylenediamine tetraacetic acid; FW, fresh weight; MOPS, 3-(N-morfolino) propanesulfonic acid; NADK, NAD kinase; NDex , external NAD(P)H dehydrogenases; NDin , internal NAD(P)H dehydrogenases; PVP, polyvinylpyrrolidone; SHAM, salicylhydroxamic acid; TES, 2-[[1,3-dihydroxy2-(hydroxymethyl)propan-2-yl]amino]ethanesulfonic acid; WWC, weight water content. ∗ Corresponding author. Tel.: +48 22 5543008; fax: +48 22 5542022. E-mail address: [email protected] (I.M. Juszczuk). 0176-1617/$ – see front matter © 2013 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.jplph.2013.12.013

SO4 2− across the plasma membrane (Hawkesford et al., 2003). ATP is constitutively required in a key reaction of the S assimilation pathway that is catalyzed by ATP-sulphurylase, which mediates the synthesis of adenosine 5 -phosphosulphate using SO4 2− . Thus, ATP availability determines SO4 2− adenylation and production of metabolically useful sulphide (Yi et al., 2010). In non-photosynthetic tissues such as roots, ATP is generated mainly in the mitochondria, whereas in the leaves, ATP generated in the chloroplast electron transport chain is supplemented by mitochondrial synthesized ATP (Hoefnagel et al., 1998). Sulphate is reduced to sulphide in plastids, but cysteine synthesis occurs ubiquitously in the plastids, cytosol, and mitochondria (Hell and Wirtz, 2011). Thus, in terms of energy provision, S metabolism largely depends on mitochondrial activity. Furthermore, inside the plant mitochondria, S is utilized for the assembly of the iron–sulphur (Fe–S) clusters. This prosthetic group in proteins contains acid-labile sulphide attached to Fe, which is essential for respiration, photosynthesis,

550

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

and nitrogen fixation (Balk and Pilon, 2011). Proteins of the mitochondrial respiratory chain contain several Fe–S clusters located in Complex I (Ohnishi, 1998) and in Complexes II and III (Balk and Pilon, 2011). Moreover, cysteine participates in disulphide bridge formation. Activity of alternative oxidase (AOX), a mitochondrial respiratory chain enzyme, is dependent on the redox state of two cysteine residues (Albury et al., 2009). Considering these functions of S, S accessibility is essential for proper mitochondrial functioning. A previous report demonstrated that complete lack of S highly influences mitochondrial respiratory chain activity in bean plants (Juszczuk and Ostaszewska, 2011). Plants exhibit acclimation to stress by increasing their metabolic resistance, and mitochondria play a pivotal role in this process (Suzuki et al., 2012). Plant mitochondria have a highly branched respiratory chain, due to the occurrence of the additional internal and external type II NAD(P)H dehydrogenases (NDin/ex ) and AOX (Rasmusson et al., 2004). These additional respiratory chain components allow flexibility of mitochondrial function in response to a variable environment (Rasmusson et al., 2009). Complex I and the cytochrome chain activities can be readily unbalanced by fluctuations in nutrient availability, such as phosphate (P), Fe, or S; however, at times of nutrient deficiency, the additional enzymes of the branched chain can partially sustain respiration (Rychter and Mikulska, 1990; Juszczuk et al., 2007; Vigani et al., 2009; Juszczuk and Ostaszewska, 2011; Vigani, 2012). Plant mitochondria are characterized by their multifunctional role, biochemical plasticity, and dynamic morphology (Logan, 2006). Few data are available that correlate the functioning of mitochondria with their ultrastructure in response to mineral starvation. There are some reports concerning P and Fe deficiencies only (Wanke et al., 1998; Vigani et al., 2009). Biochemical data in combination with transmission electron microscopy (TEM) have been invaluable for studying plant adaptations to insufficient nutrient supply; however, such a combinatorial examination of mitochondria has been reported rarely. The majority of studies of the metabolic modifications in response to S deficiency have been performed using the model plant, Arabidopsis thaliana. Transcriptomic and metabolomic analyses uncovered a connection between S nutrition and different carbon/nitrogen (N)/S metabolic pathways, including energy transformation reactions during photosynthesis (Nikiforova et al., 2005). Mitochondria participate in the adaptation response to nutritional starvation, because the process requires the reorganization of primary metabolism owing to the need for a constant supply of NAD(P)H and ATP. However, to our knowledge, this is the first report of the influence of S deficiency on the functioning and ultrastructure of mitochondria. The purpose of our study was to investigate changes that mitochondria undergo and the role of the branched electron transport chain in response to S deficiency.

Materials and methods

Co(NO3 )2 ·6H2 O, 159 nM Al2 (SO4 )3 , 1.77 ␮M MnCl2 ·4H2 O, 221 nM NiSO4 ·4H2 O, 150 nM KI, 210 nM KBr, 207 nM Na2 MoO4 ·2H2 O. The S-deprived plants were grown in nutrient solutions like the control but with SO4 2− salts replaced with comparable chloride salts in equal concentrations (S-deficient). The nutrient solution was changed twice a week. Plants were grown in a growth chamber at 150 ␮mol m−2 s−1 photosynthetically active radiation (daylight and warm white 1:1; LF-40W; Piła, Poland) with an 8 h light/16 h dark cycle at 22 ◦ C/19 ◦ C, respectively, and 60–70% relative humidity. After 9 weeks of growth in the nutrient media, leaves or roots from plants were collected for the assays in the 4th h of the light period.

Determination of tissue weight water content and biomass parameters For each treatment, 6 to 50 plants were randomly selected. Fresh tissue material was weighed (fresh weight; FW), and the same plants were used for biomass determination after oven-drying at 55 ◦ C for 4 d (dry weight; DW). Then, the tissue weight water content (WWC) was calculated as: WWC (%) = 100 × [(FW − DW)/(FW)]. To measure the surface area of all leaf blades from an individual rosette, leaves were placed under a translucent, anti-reflective, fold-up cover that presses and holds leaves flat, and the area was measured using WinFOLIAPro software program (Regent Instruments Inc., Canada). The number of leaves in a selected rosette was also determined. Root length was measured after removing the remaining nutrient medium from the roots by blotting with paper towels. Soluble protein concentration was determined by Bradford’s assay (1976) using bovine serum albumin (BSA) as a standard.

Mineral concentration analysis S, SO4 2− , and selected macro- and micronutrients were estimated in oven-dried leaf or root tissue samples. Roots were washed at 4 ◦ C in distilled water for 5 min, desorbed in 5 mM CaCl2 for 10 min, to remove unbound and weakly bound metal from the apoplast, and again washed in water for 10 min. To determine total S and macro- and micronutrient concentrations, 1 g of plant tissue was mineralized in a mixture of HNO3 and HClO4 (4:1, w/w) during 1.5 h (first 30 min at 80 ◦ C and then 1 h at 120 ◦ C). Sulphate was estimated after the digestion of 5 g of plant tissue in 50 mL of 2% CH3 COOH and 1 h of systematic vortexing. Elemental contents were determined by inductively coupled plasma optical emission spectrometry (Thermo Elemental-IRIS Advantage, USA). The analyses were performed at Warsaw University of Life Sciences, in cooperation with an authorized Laboratory of Forest Environment Chemistry of the Forest Research Institute, Poland (No AB 740 according to Polish Centre for Accreditation).

Plant material and growth conditions Arabidopsis thaliana ecotype Col-0 seeds were germinated in seed holders containing 1% (w/v) sterile plant agar. Excess seedlings were removed 7–10 d after sowing, leaving one seedling of similar size per holder. Plants were grown in a hydroponic growing system (Araponics SA, Belgium) using 2 L containers, which were filled with distilled water, kept covered with a transparent lid for 1 week, and then filled with the modified Knop nutrient medium flushed with air. The complete nutrient solution (Control) included the following components: 3 mM Ca(NO3 )2 , 1.5 mM KNO3 , 1.25 mM MgSO4 , 1 mM KH2 PO4 , 0.04 mM FeEDTA, 200 nM CuSO4 ·5H2 O, 348 nM ZnSO4 ·7H2 O, 5.6 ␮M H3 PO4 , 172 nM

Extraction and determination of pigment concentrations Chlorophylls a and b and carotenoids were extracted from 80 mg of leaf tissue ground in liquid N2 with 1.5 mL of 80% (v/v) acetone. The extract was centrifuged at 10,000 × g for 10 min. The supernatant was separated, and the absorbance values were measured at 663.2 nm, 646.8 nm, and 470 nm on a Shimadzu UV-260 spectrophotometer (Shimadzu, Japan). All absorbance values were normalized by subtracting the absorbance at 750 nm. The amount of chloroplastic pigments was calculated according to the formulas of Lichtenthaler (1987).

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

Measurements of starch concentrations For starch extraction, approximately 0.25 g FW of leaf and approximately 0.5 g FW of root tissue were weighed and ground in a pre-cooled mixer-mill (MM 200, Retsch, Germany) with 1.5 mL of 80% (v/v) ethanol. Soluble sugars were extracted by heating the samples at 37 ◦ C for 2 h. Homogenates were clarified by centrifugation at 10,000 × g for 10 min. The extractions were repeated twice, and the pellet was analyzed for starch content. Pellets were resuspended in 3 mL of water and boiled for 3 h. Amyloglucosidase (Sigma–Aldrich; cat. no. A7095) was used to digest samples overnight. Released glucose was determined spectrophotometrically by a glucose oxidase-peroxidase reaction (Kunst et al., 1985). Isolation of mitochondria Due to the lower isolation efficiency of intact mitochondria from S-deficient plants, different amounts of tissue were required from control and stressed plants; mitochondria were isolated at 4 ◦ C from 60 g of control and 80 g of S-deficient leaves and from 15 g control and 25 g S-deficient roots. Plant tissue was ground, using a cold mortar and pestle with a small amount of quartz (fine granular, washed and calcined for analysis; Merck), in 500 mL (leaves) or 250 mL (roots) of homogenization medium containing the following components: 0.45 M mannitol; 60 mM 2-[[1,3-dihydroxy2-(hydroxymethyl)propan-2-yl]amino]ethanesulfonic acid (TES), pH 8.0; 10 mM EDTA; 10 mM KH2 PO4 ; 25 mM Na4 P2 O7 ; 1 mM glycine; 1% (w/v) polyvinylpyrrolidone 40 (PVP-40); and 1% (w/v) BSA. The homogenate was filtered through an 80-␮m nylon mesh, and mitochondria were isolated by differential centrifugations: 2500 × g for 5 min and 15,000 × g for 15 min (Eppendorf 5804R centrifuge, rotor F-34-6-38). The pellet obtained was resuspended and centrifuged again at 2500 × g for 5 min and 15,000 × g for 15 min. The resulting pellet was resuspended in 6 mL of the homogenization medium, and mitochondria were purified on a discontinuous Percoll gradient using a modified method of Nishimura et al. (1982). The discontinuous gradient was prepared, stored at 4 ◦ C until used, and composed of 4.5 mL 60% (v/v) Percoll, 6 mL 45% (v/v) Percoll, 12 mL (leaves) or 6 mL (roots), 28% (v/v) Percoll, and 6 mL 10% (v/v) Percoll, in the following mixture: 0.45 M mannitol; 20 mM MOPS, pH 7.2; 1% (w/v) PVP-25; and 1% (w/v) BSA. The crude mitochondrial fraction was carefully layered on top of pre-formed gradients and centrifuged at 7000 × g (Beckman Avanti centrifuge, angle rotor F0650) for 1 h 40 min (leaves) or 1 h (roots). The mitochondrial fraction, characterized by high integrity, appeared at the interface between the 28% and 45% (v/v) Percoll layers. To obtain purified mitochondria, the mitochondrial fraction was collected, washed in a medium containing 0.45 M mannitol; 10 mM TES, pH 7.5; and 10 mM KH2 PO4 , and centrifuged at 15,000 × g for 20 min. The pellet was suspended in approximately 150 ␮L of washing medium. The concentrations of mitochondrial protein in micrograms per gram FW were for leaves: 7.80 ± 0.89 control and 10.19 ± 0.49 S-deficient, for roots: 42.81 ± 2.61 control and 55.99 ± 3.98 for Sdeficient. Mitochondrial integrity was determined by comparison of cytochrome c oxidase (COX, EC 1.9.3.1) activity measured in isotonic medium, in the absence and then in the presence of 0.04% (w/v) Triton X-100, to disrupt mitochondrial membranes (Wigge and Gardeström, 1987). Respiratory measurements Four leaves and one root were harvested for respiratory examination. Oxygen consumption rates were measured in an aqueous phase using a Clark-type oxygen electrode (Rank Brothers Ltd., Cambridge, UK) in the dark at a constant temperature of 25 ◦ C, with minor modifications to the method described by Florez-Sarasa

551

et al. (2009). Measurements were performed in a final volume of 3.2 mL assay solution. Tissues were weighed and pre-incubated for 20 min before measurement in an assay solution containing 30 mM MES, pH 6.2, 0.2 mM CaCl2 . For inhibitor treatment, 10 mM KCN or 20 mM salicylhydroxamic acid (SHAM) in DMSO was used. To measure residual respiration, both inhibitors were added. Oxygen uptake by isolated mitochondria was measured with a Clark-type O2 electrode (Oxygraph and Oxygraph Plus Software, Hansatech, Norfolk, UK) at 25 ◦ C. Calibrations were made from the difference in the signal between aerated water and Na-dithionite saturated water. Measurements were performed in a final volume of 0.5 mL mitochondria-containing assay buffer. The assay medium consisted of the following components: 0.45 M mannitol; 10 mM TES, pH 7.5; 10 mM KCl; 2 mM MgSO4 ; 5 mM KH2 PO4 ; 0.1% (w/v) BSA; and 100 ␮g of mitochondrial protein. Malate (10 mM; with 1 mM glutamate and 0.2 mM NAD+ ) was used as substrate to stimulate respiration. Complex I capacity was measured in the presence of 1 mM deamino-nicotinamide hypoxanthine dinucleotide (deamino-NADH Sigma–Aldrich; Johansson et al., 2004) and membrane permeabilizing alamethicin (AlaM, 50 ␮M; Gostimskaya et al., 2003). The capacity of additional NDex NADH with NADH as a substrate was measured after the addition of 1 mM CaCl2 . The capacity of additional NDin NADH was measured with malate as a substrate after the inhibition of Complex I by 1 ␮M rotenone. To measure state 3 respiration, 100 ␮M ADP was added. COX was inhibited with 800 ␮M KCN. AOX capacity was measured in activating conditions, in the presence of 1 mM pyruvate and 10 mM dithiothreitol, and then AOX was inhibited with 750 ␮M SHAM. Isolation of chloroplasts Plants were illuminated for 30 min at the end of the dark period, and then 25 g of leaves were collected and homogenized quickly using a cold blender (Waring) in 100 mL of the frozen grinding buffer consisting of the following components: 0.33 M sorbitol, 50 mM HEPES (pH 7.3), 0.4 mM KCl, 0.1 mM EDTA, 0.1% BSA, 0.5% PVP, and 0.2% sodium ascorbate. The homogenate was filtered through 6 layers of gauze and centrifuged at 2000 × g for 10 min at 4 ◦ C. The resulting pellet was used for the analyses following resuspension in 0.5 mL of buffer containing the following components: 0.33 M sorbitol, 50 mM HEPES (pH 7.7), 1 mM EDTA, 1 mM MgCl2 , 1 mM MnCl2 , and 1% BSA. Assays for photosystem activity Photosystem I (PSI) and photosystem II (PSII) activities in prepared chloroplasts containing 20 ␮g of chlorophyll were measured spectrophotometrically according to the method of Vernon and Cardon (1982) in HEPES buffer (pH 7.6), 330 mM sorbitol, 1 mM KH2 PO4 , 5 mM NaCl, 5 mM MgCl2 , and 5 mM NH4 Cl. Determination of adenylate concentration Concentrations of ATP and ADP were measured in leaf or root tissue extracts. Plant tissues were collected after 4 h of a light period and immediately frozen in liquid N2 . Portions of approximately 20–50 mg of frozen tissue powder were homogenized for a maximum of 3 min with 300 ␮L of 3% trichloric acid in pre-cooled Eppendorf adapters from a mixer-mill (MM 200, Retsch, Germany). The homogenate was centrifuged at 10,000 × g for 10 min at 4 ◦ C. The supernatant was stored at −20 ◦ C. ADP was converted to ATP by pyruvate kinase (Roche; cat. no. 10128155001). The ATP concentration was assayed luminometrically by the luciferin–luciferase reaction (Ludin, 2000) using a commercial ATP kit SL (cat. no. 144041, Biothema, Sweden).

552

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

Fig. 1. Effects of S deficiency on growth and morphology of 9-week-old Arabidopsis thaliana plants grown in hydroponic culture in full Knop nutrient medium (Control) or without SO4 2− (S-deficient). S-deficient plants (A), the particular rosette of a S-deficient plant (B) in comparison to control plant. Morphology of the leaves of control (C1 ) ), purple pigmentation of and S-deficient plants (C2 ). Individual leaves from control or S-deficient plants (D). Arrows indicate diagnosed chlorosis and/or necrosis ( ), and curling of the leaf lamina margins ( ). leaf stalks and veins (

Estimation of pyridine nucleotides Determination of pyridine nucleotides was performed in leaf or root tissue extracts. About 500 mg of frozen leaf powder was homogenized in a pre-cooled mortar with 2.5 mL of 0.1 M HCl (for oxidized nucleotides extraction) or 0.1 M NaOH (for reduced nucleotides extraction) in 50% ethanol. The homogenate was centrifuged at 10,000 × g for 10 min at 4 ◦ C. The acidified extracts were kept on ice. Alkaline extracts were heated at 60 ◦ C for 15 min and then cooled on ice. The supernatants were neutralized and centrifuged at 10,000 × g for 10 min at 4 ◦ C. Pyridine nucleotides in whole tissue extracts were determined by the method described by Lechevallier et al. (1977), with modifications as described by Juszczuk and Rychter (1997). Determination of NAD kinase activity Frozen leaf or root tissue powder was homogenized with 50 mM phosphate buffer (pH 7.6) and centrifuged at 15,000 × g for 15 min at 4 ◦ C. The supernatant was used for enzyme assays within 6 h. NAD kinase (EC 2.7.1.23, NADK) activity was estimated according to the method used by Hayashi et al. (2005) with modifications as described by Szal et al. (2008). Transmission electron microscopy Leaf samples were harvested from control and S-deficient plants for TEM processing. Tissues were sliced into 1–5 mm pieces and fixed in 2.5% glutaraldehyde in 0.05 M phosphate buffer at pH 7.2 for 2 h, washed in buffer, and placed for post-fixation in 2% OsO4 at 4 ◦ C in 50 mM phosphate buffer, overnight. After rinsing in phosphate buffer, the samples were dehydrated in acetone by a series of gradient washes, followed by final washes in 100% acetone. Dehydrated tissues were infiltrated with a graded series of Epon 812 resin in acetone and embedded in Epon 812 resin. The samples were polymerized at 30 ◦ C for 12 h and then at 60 ◦ C for 48 h. The tissues

were sliced into ultra-thin sections on a Leica UCT ultramicrotome and stained with uranyl acetate and lead citrate. The ultrastructures of the organelles were examined and photographed with an electron microscope JEM 1400 (JEOL Co., Japan, 2008) equipped with an energy-dispersive full-range X-ray microanalysis system (EDS INCA Energy TEM, Oxford Instruments, Great Britain), a tomographic holder, and a high resolution digital camera (CCD MORADA, SiSOlympus, Germany). The studies were performed in the Laboratory of Electron Microscopy, Nencki Institute of Experimental Biology, Warsaw, Poland. All experiments were repeated four times. Statistical analyses All presented data are the mean of n replicates (n = 3 to 50) with standard deviation (±SD). The statistical significance of the data was tested using the Student’s t-test in Excel 2007 (Microsoft software). Different superscript letters (ab , cd , ef , or gh ) indicate significant differences at P < 0.05. Results A. thaliana plants subjected to continuous S starvation for 9 weeks were smaller than plants grown in control nutrient medium. The size of a particular rosette of an S-deficient plant was smaller than the control (Fig. 1A and B), and the rosettes contained fewer leaves than those of control plants (Table 1). The surface area of all the leaves in a rosette of a randomly selected plant was less in the S-deficient plant, but root length did not significantly differ, compared to that of the control (Table 1). The FW and DW of the leaves of S-deficient plants were lower than those of control plants, but the FW and DW of the roots were not statistically different; therefore, the rosette to root dry mass ratio decreased by about 30% (Table 1). The WWC in S-deficient plants did not differ either in the leaves nor in the roots, as compared to control (Table 1). Total S content was over 50% and 60% lower in the leaves and roots of S-deficient plants, respectively, as compared to control

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

553

Table 1 Effects of S deficiency on growth parameters of Arabidopsis plants. WWC, weight water content. The surface area of the leaves and the number of leaves (calculations for randomly selected rosettes). Values are means ± SD from 6 to 50 replicates. Statistically significant differences at ab P < 0.05 by Student’s t-test. Leaves

Roots

Control 3.24 ± 0.237a ± 92.69 ± 179a ± 40a ± – 9.48a a

FW (g) DW (g) WWC (%) Surface area of leaves (cm2 ) Number of leaves Roots length (cm) Rosette DW/Root DW

S-deficient 2.27 ± 0.172b ± 92.34 ± 124b ± 35b ± – 6.61b b

0.53 0.044 0.29 6 4

0.76 0.056 1.25 34 4

Control 0.48 ± 0.025 ± 94.69 ± – – 26 ±

S-deficient 0.11 0.005 0.59

4

0.41 ± 0.026 ± 93.62 ± – – 26 ±

0.09 0.006 1.15

5

Table 2 Effects of S deficiency on mineral concentration in the leaves and roots of Arabidopsis plants. ND, not determined. Values are means ± SD from 5 to 15 replicates. Statistically significant differences at ab P < 0.05 by Student’s t-test. Leaves

S (mg g−1 DW) SO4 2− (mg g−1 DW) N (mg g−1 DW) Fe (␮g g−1 DW) S/N

Roots

Control

S-deficient

7.72a ± 0.50 3.86a ± 0.31 58.90 ± 4.91 96.24 ± 14.27 0.13

3.68b ± 0.71 1.48b ± 0.20 51.09 ± 9.27 96.41 ± 11.77 0.07

(Table 2). SO4 2− concentration was about 62% lower in the leaves of S-deficient plants compared to control leaves (Table 2). S deficiency had no influence on total N concentration (Table 2). However, since S content was significantly lower in the leaves of S-deficient plants (Table 2), the S/N ratio decreased from the value of 0.13 in control leaves to 0.07 in S-deficient leaves (Table 2). Fe concentration was similar in control and in S-deficient tissues (Table 2). Young leaf blades of S-deficient plants were sometimes misshapen, showing curling of the margins and protuberances localized in the central area (Fig. 1D). The leaf stalks became visibly violet, and this pigmentation was characteristic, especially of the late stage of S starvation (Fig. 1C2 and D). Chlorotic or necrotic spots were visible on the surface of several leaves (Fig. 1D). Examination of photosynthetic pigments revealed unchanged concentrations of chlorophylls and carotenoids. The activities of PSI and PSII in chloroplasts from Sdeficient plants were both approximately 30% less than those in the control (Table 3). The ultrastructure of mitochondria and chloroplasts in Sdeficient and control leaf mesophyll tissues were examined by TEM (Figs. 2 and 3). Chloroplasts from S-deficient plants exhibited a typical ellipsoidal shape with well-arranged thylakoid membranes in distinct grana regions. The stroma contained numerous plastoglobuli; however, fewer starch grains accumulated in the chloroplastic stroma as compared to control plants (Fig. 3). Starch concentration in leaf extracts was lower for S-deficient plants compared to control, 53 ± 2 mg g−1 FW versus 75 ± 2 mg g−1 FW, respectively. The mitochondria from control plants displayed dense, granular,

Table 3 Effects of S deficiency on photosynthetic pigment (chlorophyll a and b, carotenoids) concentrations, and PSI, PSII activities in Arabidopsis leaves. Values are means ± SD from 12 replicates (photosystems) and 30 replicates (chlorophylls and carotenoids). Statistically significant differences at ab P < 0.05 by Student’s t-test. Leaves Control −1

Chlorophyll a (mg g FW) Chlorophyll b (mg g−1 FW) Carotenoids (mg g−1 FW) PSI (␮mol TMPD mg chl−1 h−1 ) PSII (␮mol DCPIP mg chl−1 h−1 )

0.82 0.30 0.13 76.87a 14.84a

± ± ± ± ±

S-deficient 0.11 0.04 0.03 6.65 2.12

0.83 0.30 0.13 55.63b 10.71b

± ± ± ± ±

0.09 0.04 0.02 7.90 1.87

Control 10.96a ± 0.05 ND ND 4638.85 ± 201.65 –

S-deficient 4.41b ± 0.23 ND ND 4945.69 ± 368.75 –

and homogeneous matrices as well as continuous, regularly folded inner membrane forming translucent cristae (Fig. 2A–C). In marked contrast, many mitochondria from S-deficient Arabidopsis exhibited a swollen morphology (control plants, 1 per image, n = 50; S-deficient plants, 3 per image, n = 47; for the images of the same magnification), although some mitochondria were not noticeably affected by S deficiency (Fig. 2D). Dilated mitochondria displayed a lower matrix density and fewer cristae than mitochondria from control plants (Fig. 2E and F). Disintegration of the inner and outer membranes of some mitochondria was observed (Fig. 2F). S deficiency resulted in the decrease in oxygen consumption both in leaves and roots by 40 and 35%, respectively, as compared to control plants (Table 4). After KCN addition the total oxygen consumption by the control and S-deficient leaves and roots decreased to the similar value, however KCN inhibited more the respiration of control tissues (about 75%) than S-deficient tissues (about 55–65%) indicating lower cytochrome pathway capacity in S-deficient leaves and roots (Table 4). The presence of SHAM, AOX inhibitor, diminished respiration by about 40% in all the tissues and the remaining O2 consumption for S-deficient leaves and roots was 38% and 31% lower as regard to control, respectively (Table 4). The residual O2 consumption rate which is the fraction of oxygen uptake that is resistant to the combination of KCN and SHAM (Ribas-Carbó et al., 1997) was 2–6% of the initial O2 consumption rates in all tissues (Table 4). Purified mitochondria isolated from the leaves and roots of control and S-deficient A. thaliana plants showed high integrity of the external membrane, as expressed by cytochrome c activity, between 80% and 90% (Table 5). The levels of respiratory control were unchanged for all substrates tested; whereas, ratios of ADP/O were lower in mitochondria isolated both from the leaves and roots of S-deficient plants (Table 5). Prolonged S starvation modified the respiratory activity of mitochondria. In mitochondria isolated from the leaves and roots of S-deficient plants, respiration with malate, which is a Complex I substrate, was 40% and 25% lower, respectively, as compared to respiration in control mitochondria. Lower Complex I capacity in S-deficient plants was confirmed by the use of deamino-NADH, a specific Complex I substrate (Johansson et al., 2004), in the presence of AlaM, a channel-forming peptide which allows the passage of low-molecular mass compounds into the mitochondrial matrix (Gostimskaya et al., 2003). Similarly as with

554

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

Fig. 2. Effects of S deficiency on the ultrastructure of mitochondria in mesophyll cells of 9-week-old Arabidopsis thaliana plants grown in hydroponic culture on full Knop nutrient medium (Control) (A–C) or without SO4 2− (S-deficient) (D–F). Mitochondria with dense matrices, granular, and homogeneous (A–D). The continuous and regularly folded inner membrane (A–D). The degradation of the mitochondrial matrix and the significant reduction of the number of mitochondrial cristae (E and F). Abbreviations: CH, chloroplast; CW, cell wall; M, mitochondrion; N, nucleus; P, plastoglobule (indicated by black arrow); V, vacuole. Bar size is 500 nm.

Fig. 3. Effects of S deficiency on the ultrastructure of chloroplasts in mesophyll cells of 9-week-old Arabidopsis thaliana plants grown in hydroponic culture on full Knop nutrient medium (Control) (A) or without SO4 2− (S-deficient) (B). S-deficient Arabidopsis plants with fewer starch grains in the stroma of chloroplasts. Abbreviations: CH, chloroplast; CW, cell wall; M, mitochondrion; P, plastoglobule (indicated by black arrow); S, starch grain; V, vacuole. Bar size is 1 ␮m.

malate, deamino-NADH oxidation was lower in S-deficient plants than control plants, by approximately 45% in the leaves and 25% in the roots (Table 5). In the leaves and roots of control plants, the addition of a Complex I inhibitor, rotenone, strongly diminished oxygen consumption with malate (40–50% inhibition), but in the tissues of S-deficient plants, rotenone inhibited oxygen consumption by only 10–20%. The remaining oxygen consumption, not inhibited by rotenone, was similar in mitochondria from control and S-deficient tissues, indicating that there were no differences in the NDin NADH capacity (Table 5). Compared to control plants, the capacity of NDex NADH (measured with NADH in the presence of calcium ions) in S-deficient plants increased by greater than 45% in leaf mitochondria and by approximately 20% in root mitochondria (Table 5). COX capacity decreased by approximately 37% and 30% in the S-deficient leaf and root mitochondria, respectively (Table 5),

but the AOX capacity did not change with any substrates tested (Table 5). S deficiency modified the energy status in the tissues of Arabidopsis plants. The total concentrations of adenylates (expressed as ATP + ADP) measured in the light, were lower in the leaves and roots of S-deficient plants as compared to control, mainly due to the severe decrease of ATP level by greater than 30% (Fig. 4A and B). As a consequence of lower [ATP] and higher [ADP] in the leaves of S-deficient plants, the ATP/ADP ratio was halved, as compared to control (Fig. 4A). In the roots of S-deficient plants, [ATP], [ADP], and the ratio of ATP/ADP declined (Fig. 4B). The total [NAD(H)] decreased in S-deficient leaves, while [NAD(H)] in the roots remained unchanged, compared to control tissues (Fig. 5A and B). However, the redox level almost doubled in both leaves and roots of S-deficient plants (Fig. 5A and B, insets).

Table 4 Effects of S deficiency on respiratory activity of intact leaves and roots of Arabidopsis plants. Inhibitor concentrations were 10 mM KCN and 20 mM SHAM. Values are means ± SD from 2 independent experiments (assayed in triplicate). Statistically significant differences at ab P < 0.05 by Student’s t-test. Leaves Control Oxygen consumption rate (nmol O2 g−1 FW min−1 ) 61.92a ± 4.23 Initial rate 17.06 ± 1.49 + KCN 36.31a ± 3.55 + SHAM 2.07 ± 0.19 + KCN + SHAM

Roots S-deficient 37.50b 17.51 22.55b 2.24

± ± ± ±

3.21 1.83 1.60 0.34

Control 81.89a 18.70 47.39a 1.77

± ± ± ±

S-deficient 5.86 0.92 3.26 0.50

53.35b 17.90 32.68b 2.43

± ± ± ±

4.95 1.51 2.97 0.18

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

555

Table 5 Effects of S deficiency on respiratory activity of mitochondria isolated from the leaves and roots of Arabidopsis plants. Substrate concentrations were 10 mM malate (with 2 mM glutamate and 0.5 mM NAD+ ), 1 mM NADH (with 1 mM CaCl2 ), 1 mM deamino-NADH, and 0.08 mM ADP. Mitochondrial protein (100 ␮g) was used for the respiratory measurements. *In AOX-activated conditions. Values are means ± SD from 3 to 6 independent mitochondrial isolations. Statistically significant differences at ab P < 0.05 by Student’s t-test. Leaves

Roots

Control Oxygen consumption rate (nmol O2 mg−1 protein min−1 ) Malate Malate + rotenone Deamino-NADH NADH Respiratory control Malate NADH ADP/O Malate NADH AOX capacity* (nmol O2 mg−1 protein min−1 ) Malate NADH COX capacity (nmol cyt c mg−1 protein min−1 ) Integrity of mitochondrial membranes (%)

54a 27 33a 40b

± ± ± ±

S-deficient 33b 21 18b 59a

7 3 6 4

24b 19 23b 36a

± ± ± ±

3 1 5 3

1.2 ± 0.2 1.2 ± 0.1

2.1a ± 0.1 1.9 ± 0.2

1.5b ± 0.3 1.6 ± 0.2

2.3a ± 0.2 1.7 ± 0.3

1.7b ± 0.4 1.4 ± 0.1

17 10 750a 91

± ± ± ±

2 2 143 2

16 11 474b 90

Control e 3.24 ± 0.13

S-deficient f 1.67 ± 0.12

± ± ± ±

1 3 182 2

20 13 919a 89

± ± ± ±

5 1 250 3

15 14 647b 87

± ± ± ±

1 2 156 4

which is contrary to previous findings by López-Bucio et al. (2003), Nikiforova et al. (2003), and Zhang et al. (2011). These differences may result from a shorter light period and natural variations in Arabidopsis ecotypes or allelic-specific stress responses (Buescher et al., 2010; Prinzenberg et al., 2010; Trontin et al., 2011). The analysis of the ultrastructure of S-deficient Arabidopsis leaf mesophyll cells showed that many mitochondria became swollen and had disintegrated cristae (Fig. 2E and F) and some had disrupted outer membranes (Fig. 2F). These observations are the first ones concerning the influence of S deficiency stress on mitochondrial ultrastructure. The effect of drought stress on mitochondrial structure has been reported for poplar (Zhang et al., 2010) and apple (Wang et al., 2012). Those studies show that mitochondria from control plants possessed extensive cristae appearing as irregularly oriented sacs and folds; in marked contrast, drought stressed plants had swollen mitochondria, and the cristae disappeared. Ultrastructure analysis of Fe-deficient cucumber root mitochondria revealed typical elongations in the central parts of the mitochondria (Vigani et al., 2009). The internal mitochondrial membrane contains all the protein complexes organized in the mitochondrial respiratory chain. Thus, lack of continuity of the internal mitochondrial membrane observed in some mitochondria from S-deficient leaves (Fig. 2E and F) disturbs electron flux and may lead to an energy deficiency in Arabidopsis S-deficient plants (Fig. 4). An effect of S deficiency

B

40

ATP/ADP

Control 8.40e ± 0.56

S-deficient 6.57f ± 0.34

40 bc

30

35 30

20

ADP

15

ATP

25 bd

20

10

5

5

ADP ATP

15

10

0

ac

-1

25

nmol g FW

ad

-1

nmol g FW

2 2 1 1

1.6 ± 0.2 1.2 ± 0.2

A. thaliana plants showed morphological symptoms of S deficiency (Fig. 1 and Table 1), changes in the respiratory activity (Tables 4 and 5), and modifications of cellular energy and redox homeostasis (Figs. 4 and 5). The decreases in total S and SO4 2− concentrations in the leaf and root tissues of S-deficient plants and lower S/N ratio, together with unchanged Fe concentration (Table 2) further support the conclusion that the exhibited symptoms of stress were specific to S deficiency. All stress symptoms of S deficiency appeared after 7 weeks of Arabidopsis cultivation in an 8-h photoperiod. Nikiforova et al. (2003) indicate the appearance of S stress symptoms earlier, after only 11 d of plant growth in a 16-h photoperiod. Moreover, we did not observe differences in the root length of Arabidopsis S-deficient and control plants (Table 1),

35

± ± ± ±

1.4 ± 0.3 1.3 ± 0.1

Discussion

ATP/ADP

32a 19 30a 30b

5 6 1 7

S-deficient

1.8 ± 0.6 1.5 ± 0.2

A high NADH/NAD ratio in S-deficient tissues resulted from the 45% higher [NADH] and decreased [NAD] (25% in leaves and 10% in roots), compared to control plants (Fig. 5A and B). S deficiency resulted in the increase in total NADP(H) level in the leaves because of 45% higher [NADP], as compared to control (Fig. 5C). In the roots of S-deficient plants, total [NADP(H)] did not change significantly, and the NADPH/NADP ratio only slightly decreased (Fig. 5D, inset). Estimated NADK activity almost doubled in S-deficient leaf tissue (Fig. 5C, inset), compared to control NADK activity.

A

± ± ± ±

Control

0

Control

S-deficient

Control

S-deficient

Fig. 4. Effects of sulphur deficiency on [ATP] and [ADP] in the leaves (A) and roots (B) of Arabidopsis plants. Values are means ± SD from 6 to 9 replicates. Statistically significant differences at ab,cd,ef P < 0.05 by Student’s t-test. Letters ab correspond to ADP, cd to ATP, and ef to ATP/ADP ratio values in the table insets on the graphs.

556

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

NADH/NAD

120

S-deficient e 0.20

B

NADH/NAD

NADH

60

NAD

40

80 NADH

60 40

20

b

a

NAD

20

0

0

Control S-deficient

C NADPH/NADP NADK

Control e 0.79 456h ± 65

Control S-deficient S-deficient f 0.48 873g ± 62

D 60

c

40 NADPH

30

NADP

20

NADPH/NADP

Control 0.76e

S-deficient 0.68f

50

d

10

nmol g-1 FW

50

nmol g-1 FW

S-deficient e 0.22

100

ad

80

60

Control f 0.14

120

bc

100

nmol g-1 FW

Control f 0.10

nmol g-1 FW

A

40 NADPH

30

d

c

20

NADP

10

0

0

Control S-deficient

Control S-deficient

Fig. 5. Effects of S deficiency on pyridine nucleotide concentrations and NADK activity in the leaves (A, C) and roots (B, D) of Arabidopsis plants. Values are means ± SD from 6 to 9 replicates. In the tables without SD values, SD does not exceed 2%. Statistically significant differences at ab,cd,ef,gh P < 0.05 by Student’s t-test. Letters ab correspond to NAD(P)H, cd to NAD(P), ef to NAD(P)H/NAD(P) ratio, and gh to NADK activity (nmol NADP g−1 FW h−1 ) values in the table insets on the graphs.

is a general impairment of respiratory activity, which has been observed on the level of whole tissues (Table 4) and isolated mitochondria (Table 5). The total respiratory rates and cyanidesensitive respiration of the leaves and roots of S-deficient plants were always lower as compared to control, and this decrease concerned cyanide-sensitive respiration, whereas cyanide-resistant and SHAM-sensitive respiration remained similar in both control and S-deficient plants (Table 4). The Percoll purification procedure to isolate mitochondria eliminates organelles having discontinuous outer membranes and allows for isolation of only high integrity mitochondria, which may be used for the respiratory chain activity assay. In mitochondria isolated from S-deficient leaves and roots, oxygen consumption with malate as a substrate was diminished and was accompanied by decreased ADP/O ratios which may be the result of lower Complex I activity (Table 5). A decreased respiratory capacity of Complex I was also demonstrated by rotenone-inhibited malate oxidation and lower oxygen consumption with an NADH analogue, deamino-NADH (Table 5), which is readily oxidized only by Complex I (Johansson et al., 2004). Complex I contains most of the Fe–S clusters in the respiratory chain (Lin et al., 1995). Thus, the decrease in its activity could indicate a lower amount of protein, improper structure of the polypeptide, and/or improper Fe–S cluster assembly. A decrease in Complex I capacity was observed also in Fe-deficient cucumber root mitochondria and is attributed to impaired synthesis of subunits containing Fe–S clusters (Vigani et al., 2009). Complex I and COX, which are involved in proton translocation, contribute to the proton gradient across the inner mitochondrial membrane and thus regulate ATP biosynthesis

(Dudkina et al., 2006). Strongly decreased COX capacity has been shown previously in leaf and root mitochondria of S-deficient bean (Juszczuk and Ostaszewska, 2011), Fe-deficient sycamore cells (Pascal and Douce, 1993), and in the root mitochondria of Fedeficient cucumber (Vigani et al., 2009). Decreases in Complex I and COX capacities (Table 5) partially explain the lower ATP level. A small and rapidly changing adenylate pool depends on the efficiency of processes producing and consuming ATP for growth and maintenance (Gardeström and Wigge, 1988; Mikulska et al., 1998). The level of ATP is critical for modulating S flow through its assimilation pathway (Yi et al., 2010). During S starvation, the ATP requirement likely increases due to enhanced uptake, transport, and assimilation of SO4 2− in the leaves. Increased activity of SO4 2− transporters in the root cells were found in Arabidopsis plants as a result of S depletion (Yoshimoto et al., 2007; Shinmachi et al., 2010). The increased ATP demand for S uptake and assimilation may limit other cellular use of ATP for growth processes in S-deficient Arabidopsis, resulting in the observed inhibition of plant growth (Table 1 and Fig. 1). Decreased ATP levels under S deficiency were found in the roots of broad bean (Pacyna et al., 2006), pea and alfalfa (Scherer et al., 2008), and in the leaves and roots of common bean (Juszczuk and Ostaszewska, 2011). Adenylate concentration in the extracts from light-exposed green tissues reflects the chloroplastic, mitochondrial, and cytosolic pools; whereas, adenylate concentration in the roots reflects the contribution of mitochondrial and cytosolic ATP pools only. The decrease in total adenylates was more pronounced in the roots (32%) than in the leaves (16%) (Fig. 4), but the decrease in [ATP] was similar in Arabidopsis leaves and roots, 32% and 35%, respectively

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

(Fig. 4). Additionally, in the leaves, the decrease in [ATP] was accompanied by increased [ADP] (Fig. 4). We did not estimate adenylate concentration separately in the chloroplasts, but the decreased [ATP] and increased [ADP] in the leaves (Fig. 4) could be explained by lower ATP synthesis in the chloroplasts. In S-deficient Arabidopsis leaves, lower chloroplast ATP synthase activity was observed due to decreased mRNA of atpA, atpB, atpE, and atpF (Kandlbinder et al., 2004). In the roots, the more pronounced decrease in total adenylates (Fig. 4) could be the direct consequence of lower mitochondrial contribution. We did not observe any effect of S deficiency on chlorophyll a and b or carotenoid concentrations (Table 3), leaf NADPH levels (Fig. 5B), or chloroplast ultrastructure (Fig. 3). However, maximum activities of PSI and PSII were approximately 30% lower than those in the control (Table 3), fewer starch grains accumulated in the stroma of chloroplasts (Fig. 3), and starch concentration in the leaves was lower in the S-deficient plants than in the controls. Further, notable reductions of rosette biomass (Table 1) and ATP levels (Fig. 4) indicate energy deficiency. Unchanged chlorophyll content and photosynthetic yield for PSII were shown previously in A. thaliana under short-term S deficiency (Kandlbinder et al., 2004). Other investigations indicate lower levels of chlorophylls in the leaves of S-deficient tomato (Zuchi et al., 2009), mulberry (Tewari et al., 2010), and Arabidopsis (Nikiforova et al., 2005). Wulff-Zottele et al. (2010) report that, in Arabidopsis, chlorophyll content decreased only under short S depletion combined with high light intensities. In our experiments, S deficiency in Arabidopsis plants had no influence on NADP(H) level in the roots; whereas in the leaves, the NADPH/NADP ratio decreased markedly, owing to increased [NADP] (Fig. 5). NADPH oxidation likely exceeded NADP reduction by the electron carriers of the photosynthetic electron transport chain, because of the lower activity of PSI and PSII (Fig. 2). Moreover, the increased NADP(H) level accompanied by the decreased NAD(H) level can be explained by the elevated NADK activity, which was almost doubled in the tissues of S-deficient plants compared to control plants (Fig. 5C). NADK catalyses the phosphorylation of NAD+ to form NADP+ , using ATP as a phosphoryl donor, and has an important function in supplying NADP+ /NADPH as well as in regulating the ratio of NAD(H)/NADP(H) (Shi et al., 2009). In the leaves of S-deficient Arabidopsis, a two-fold increase in NADK activity (Fig. 5C) caused 23% lower NAD levels (Fig. 5A) and 45% higher NADP levels (Fig. 5C). NADK activity is essential because NADPH supports reductive biosynthetic reactions and defends cells against oxidative stress (Grose et al., 2006; Shi et al., 2009). Similar to the previous observations in bean plants subjected to cold shock (Ruiz et al., 2002), the activity of NADK increased (Fig. 5) in the leaves of S-deficient Arabidopsis plants, thereby resulting in the higher level of NADP(H) at the expense of NAD(H) (Fig. 5). Respiratory chain NDex NADH regulates the reduction of the cytosolic pool of NADH (Moore et al., 2003; Rasmusson et al., 2004). Under S deficiency, the NADH/NAD ratio almost doubled in both leaves and roots of Arabidopsis plants (Fig. 5). Disturbances in cell redox homeostasis have been observed under different stress conditions (Rasmusson and Wallström, 2010). NADH turnover is crucial for cell metabolism, and due to presence of additional NAD(P)H dehydrogenases, mitochondria play an important role in maintaining cell redox homeostasis (Noctor et al., 2006; Rasmusson and Wallström, 2010). Increased activity of NDex NADH, induced in S-deficient plants (Table 5), may moderate the redox level of the cells and partially compensate for dysfunctional Complex I. An increase in external NADH oxidation has been observed in cytoplasmic male sterile mutant II, Complex I mutants of Nicotiana sylvestris (Gutierres et al., 1997), cucumber mosaic mutant 16 with decreased Complex I activity (Juszczuk et al., 2007), and Fe-deficient cucumber (Vigani and Zocchi, 2010). In beans, S deficiency resulted in the unchanged capacity of NDex NADH but increased capacity of

557

NDin NADH (Juszczuk and Ostaszewska, 2011). In Arabidopsis, S starvation had no influence on NDin NADH capacity (Table 5). Under Fe deficiency in cucumber root mitochondria, NDin capacity was only slightly stimulated compared to a large increase in NDex capacity (Vigani and Zocchi, 2010). Complex I is usually down-regulated by environmental fluctuations, while NDin bypassing Complex I are up-regulated (Rasmusson et al., 2009). Additional NDin/ex do not contain Fe–S clusters (Rasmusson et al., 2008), and fewer S atoms are required for their assembly. In Arabidopsis under S deficiency conditions, S availability might be the limiting factor for Complex I formation, or Complex I formation might be down-regulated to minimize S utilization, and thus type II NDin and NDex could sustain mitochondrial respiration. In mitochondria of S-deficient Arabidopsis plants, the capacity of AOX did not change as compared to control (Table 5). AOX does not contain Fe–S clusters, but AOX activity depends on cysteine residues (Albury et al., 2009). Thus, AOX biosynthesis could be preferentially supported during S deficiency conditions by redirecting the available S to this process. In S-deficient Arabidopsis, NDex NADH and AOX may provide an alternate pathway for oxidation of the excess reducing power (Fig. 5) in conditions where Complex I and IV are suppressed (Table 5). In response to S starvation, A. thaliana plants manifest modifications of cellular energy and redox homeostasis. Considering all the ultrastructural and biochemical changes, we can conclude that mitochondria from leaf and root cells are the organelles strongly affected by S deficiency. Although mitochondria in the cells of Sdeficient leaves manifested the damage, the enhanced plasticity of the respiratory chain probably allowed plants to survive and grow, despite the stress of S deficiency. Acknowledgements We would like to express our appreciation to Dr. Wiesław Szulc (Warsaw University of Life Sciences, Laboratory of Forest Environment Chemistry of Forest Research Institute, Warsaw, Poland) for mineral concentration analysis. We also wish to thank the reviewers for useful comments. This research work was supported by Grant N N303 800240 from the National Science Centre (NCN), Poland, given to I.M.J. and the intramural grant DSM501/86-102342 from MSHE through the Faculty of Biology, University of Warsaw, given to M.O. References Albury MS, Elliott C, Moore AL. Towards a structural elucidation of the alternative oxidase in plants. Physiol Plant 2009;137(4):316–27. Balk J, Pilon M. Ancient and essential: the assembly of iron–sulfur clusters in plants. Trends Plant Sci 2011;16:218–26. Bradford MM. A rapid and sensitive method for quantitation of microgram quantities of protein utilizing the principle of protein-dye-binding. Anal Biochem 1976;72:248–54. Buescher E, Achberger T, Amusan I, Giannini A, Ochsenfeld C, Rus A, et al. Natural genetic variation in selected populations of Arabidopsis thaliana is associated with ionomic differences. PLoS One 2010;5(6):e11081. Dudkina NV, Heinemeyer J, Sunderhaus S, Boekema EJ, Braun HP. Respiratory chain supercomplexes in the plant mitochondrial membrane. Trends Plant Sci 2006;11(5):232–40. Florez-Sarasa I, Ostaszewska M, Galle A, Flexas J, Rychter AM, Ribas-Carbó M. Changes of alternative oxidase activity, capacity and protein content in leaves of Cucumis sativus wild type and MSC16 mutant grown under different light intensities. Physiol Plant 2009;137(4):419–26. Gardeström P, Wigge B. Influence of photorespiration on ATP/ADP ratios in the chloroplasts, mitochondria, and cytosol, studied by rapid fractionation of barley (Hordeum vulgare) protoplasts. Plant Physiol 1988;88(1):69–76. Gostimskaya IS, Grivennikova VG, Zharova TV, Bakeeva LE, Vinogradov AD. In situ assay of the intramitochondrial enzymes: use of alamethicin for permeabilization of mitochondria. Anal Biochem 2003;313:46–52. Grose JH, Joss L, Velick SF, Roth JR. Evidence that feedback inhibition of NAD kinase controls responses to oxidative stress. Proc Natl Acad Sci USA 2006;103(20):7601–6.

558

M. Ostaszewska et al. / Journal of Plant Physiology 171 (2014) 549–558

Gutierres S, Sabar M, Lelandais C, Chetrit P, Diolez P, Degand H, et al. Lack of mitochondrial and nuclear-encoded subunits of complex I and alteration of the respiratory chain in Nicotiana sylvestris mitochondrial deletion mutants. Proc Natl Acad Sci USA 1997;94:3436–41. Hawkesford MJ, Buchner P, Hopkins L, Howarth JR. The plant sulfate transporter family: specialized functions and integration with whole plant nutrition. In: Davidian JC, Grill D, DeKok LJ, Stulen I, Hawkesford MJ, Schnug E, Rennenberg H, editors. Sulfur transport and assimilation in plants: regulation, interaction and signalling. Leiden: Backhuys Publishers; 2003. p. 1–10. Hayashi M, Takahashi H, Tamura K, Huang J, Yu LH, Kawai-Yamada M, et al. Enhanced dihydroflavonol-4-reductase activity and NAD homeostasis leading to cell death tolerance in transgenic rice. Proc Natl Acad Sci USA 2005;102:7020–5. Hell R, Wirtz M. Molecular biology, biochemistry and cellular physiology of cysteine metabolism in Arabidopsis thaliana. Arabidopsis Book 2011;9:e0154. Hoefnagel MHN, Atkin O, Wiskich JT. Interdependence between chloroplasts and mitochondria in the light and the dark. Biochim Biophys Acta 1998;1366:235–55. Johansson FI, Michalecka AM, Møller IM, Rasmusson AG. Oxidation and reduction of pyridine nucleotides in alamethicin-permeabilised plant mitochondria. Biochem J 2004;380:193–202. Juszczuk IM, Ostaszewska M. Respiratory activity, energy and redox status in sulphur-deficient bean plants. Environ Exp Bot 2011;74:245–54. Juszczuk IM, Rychter AM. The changes in pyridine nucleotide levels in leaves and roots of bean plants (Phaseolus vulgaris L.) during phosphate deficiency. J Plant Physiol 1997;151:399–404. ˛ Juszczuk IM, Flexas J, Szal B, Dabrowska Z, Ribas-Carbo M, Rychter AM. Effect of mitochondrial genome rearrangement on respiratory activity, photosynthesis, photorespiration and energy status of MSC16 cucumber (Cucumis sativus) mutant. Physiol Plant 2007;131:527–41. Kandlbinder A, Finkemeier I, Wormuth D, Hanitzsch M, Dietz KJ. The antioxidant status of photosynthesizing leaves under nutrient deficiency: redox regulation, gene expression and antioxidant activity in Arabidopsis thaliana. Physiol Plant 2004;120(1):63–73. Kunst A, Draeger B, Ziegenhorn J. Colorimertic methods with glucose oxidase and peroxidase. In: Bergmayer HU, Bergmayer J, Graßl M, editors. Methods in enzymatic analysis. Metabolites 1: carbohydrates, vol. VI, 3rd ed. Weinheim: Verlag Chemie; 1985. p. 178–85. Lechevallier D, Vermeersh J, Moneger R. Microanalyse du NADP+ et du NAD+ réduits et oxydés dans les tissus foliaires et dans les plastes isolés de Spirodéle et de Blé. 2. Méthode d’analyse des nucléotides pyridiniques de tissus végétaux. Physiol Vég 1977;15:63–93. Lewandowska M, Sirko A. Recent advances in understanding plant response to sulfur-deficiency stress. Acta Biochim Polon 2008;55(3):457–71. Lichtenthaler HK. Chlorophylls and carotenoids: pigments of photosynthetic biomembranes. In: Packer L, Douce R, editors. Methods in enzymology. New York: Academic Press Inc.; 1987. p. 350–82. Lin TI, Sled VD, Ohnishi T, Brennicke A, Grohmann L. Analysis of the iron–sulphur clusters within the complex I (NADH:ubiquinone oxidoreductase) isolated from potato tuber mitochondria. Eur J Biochem 1995;230:1032–6. Logan DC. The mitochondrial compartment. J Exp Bot 2006;57(6):1225–43. López-Bucio J, Cruz-Ramírez A, Herrera-Estrella L. The role of nutrient availability in regulating root architecture. Curr Opin Plant Biol 2003;6(3):280–7. Ludin A. Use of firefly luciferase in ATP-related assays of biomass, enzymes, and metabolites. Method Enzymol 2000;305:346–70. Mikulska M, Bomsel JL, Rychter AM. The influence of phosphate deficiency on photosynthesis, respiration and adenine nucleotide pool in bean leaves. Photosynthetica 1998;35(1):79–88. Moore CS, Cook-Johnson RJ, Rudhe C, Whelan J, Day DA, Wiskich JT, Soole KL. Identification of AtNDI1, an internal non-phosphorylating NAD(P)H dehydrogenase in Arabidopsis mitochondria. Plant Physiol 2003;133:1968–78. Nikiforova VJ, Freitag J, Kempa S, Adamik M, Hesse H, Hoefgen R. Transcriptome analysis of sulfur depletion in Arabidopsis thaliana: interlacing of biosynthetic pathways provides response specificity. Plant J 2003;33:633–50. Nikiforova VJ, Kopka J, Tolstikov V, Fiehn O, Hopkins L, Hawkesford MJ, et al. System rebalancing of metabolism in response to sulphur deprivation, as related by metabolome analysis of Arabidopsis plants. Plant Physiol 2005;138: 304–18. Nishimura M, Douce R, Akazawa T. Isolation and characterization of metabolically competent mitochondria from spinach leaf protoplasts. Plant Physiol 1982;69:916–20. Noctor G, Queval G, Gakière B. NAD(P) synthesis and pyridine nucleotide cycling in plants and their potential importance in stress conditions. J Exp Bot 2006;57(8):1603–20. Ohnishi T. Iron–sulfur clusters/semiquinones in complex I. Biochim Biophys Acta 1998;1364:186–206. Pacyna S, Schulz M, Scherer HW. Influence of sulphur supply on glucose and ATP concentrations of inoculated broad beans (Vicia faba minor L.). Biol Fert Soils 2006;42:324–9.

Pascal N, Douce R. Effect of iron deficiency on the respiration of sycamore (Acer pseudoplatanus L.) cells. Plant Physiol 1993;103:1329–38. Prinzenberg AE, Barbier H, Salt DE, Stich B, Reymond M. Relationships between growth, growth response to nutrient supply, and ion content using a recombinant inbred line population in Arabidopsis. Plant Physiol 2010;154(3):1361–71. Rasmusson AG, Wallström SV. Involvement of mitochondria in the control of plant cell NAD(P)H reduction levels. Biochem Soc Trans 2010;38(2):661–6. Rasmusson AG, Geisler DA, Møller IM. The multiplicity of dehydrogenases in the electron transport chain of plant mitochondria. Mitochondrion 2008;8:47–60. Rasmusson AG, Fernie AR, van Dongen JT. Alternative oxidase: a defence against metabolic fluctuations? Physiol Plant 2009;137:371–82. Rasmusson AG, Soole KL, Elthon TE. Alternative NAD(P)H dehydrogenases of plant mitochondria. Annu Rev Plant Biol 2004;55:23–39. Ribas-Carbó M, Lennon AM, Robinson SA, Giles L, Berry JA, Siedow JN. The regulation of electron partitioning between the cytochrome and alternative pathways in soybean cotyledon and root mitochondria. Plant Physiol 1997;113:903–11. Ruiz JM, Sánchez E, García PC, López-Lefebre LR, Rivero RM, Romero L. Proline metabolism and NAD kinase activity in greenbean plants subjected to coldshock. Phytochemistry 2002;59(5):473–8. Rychter AM, Mikulska M. The relationship between phosphate status and cyanideresistant respiration in bean roots. Physiol Plant 1990;79(4):663–7. Scherer HW, Pacyna S, Spoth KR, Schulz M. Low levels of ferredoxin, ATP and leghemoglobin contribute to limited N2 fixation of peas (Pisum sativum L.) and alfalfa (Medicago sativa L.) under S deficiency conditions. Biol Fert Soils 2008;44:909–16. Shi F, Li Y, Li Y, Wang X. Molecular properties, functions, and potential applications of NAD kinases. Acta Biochim Biophys Sin 2009:352–61. Shinmachi F, Buchner P, Stroud JL, Parmar S, Zhao FJ, McGrath SP, Hawkesford MJ. Influence of sulfur deficiency on the expression of specific sulfate transporters and the distribution of sulfur, selenium, and molybdenum in wheat. Plant Physiol 2010;153(1):327–36. Suzuki N, Koussevitzky S, Mittler R, Miller G. ROS and redox signalling in the response of plants to abiotic stress. Plant Cell Environ 2012;35(2):259–70. ˛ Z, Malmberg G, Gardeström P, Rychter AM. Changes in energy Szal B, Dabrowska status of leaf cells as a consequence of mitochondrial genome rearrangement. Planta 2008;227:697–706. Tewari RK, Kumar P, Sharma PN. Morphology and oxidative physiology of sulphurdeficient mulberry plants. Environ Exp Bot 2010;68:301–8. Trontin C, Tisné S, Bach L, Loudet O. What does Arabidopsis natural variation teach us (and does not teach us) about adaptation in plants? Curr Opin Plant Biol 2011;14(3):225–31. Vernon LP, Cardon S. Direct spectrophotometric measurement of photosystem I and photosystem II activities of photosynthetic membrane preparations from Cyanophora paradoxa, Phormidium laminosum, and spinach. Plant Physiol 1982;70:442–5. Vigani G. Discovering the role of mitochondria in the iron deficiency-induced metabolic responses of plants. J Plant Physiol 2012;169(1):1–11. Vigani G, Zocchi G. Effect of Fe deficiency on mitochondrial alternative NAD(P)H dehydrogenases in cucumber roots. J Plant Physiol 2010;167:666–9. Vigani G, Maffi D, Zocchi G. Iron availability affects the function of mitochondria in cucumber roots. New Phytol 2009;182:127–36. Wang S, Liang D, Li C, Hao Y, Ma F, Shu H. Influence of drought stress on the cellular ultrastructure and antioxidant system in leaves of drought-tolerant and drought-sensitive apple rootstocks. Plant Physiol Biochem 2012;51:81–9. Wanke M, Ciereszko I, Podbielkowska M, Rychter AM. Response to phosphate deficiency in bean (Phaseolus vulgaris L.) roots. Respiratory metabolism, sugar localization and changes in ultrastructure of bean root cells. Ann Bot 1998;82:809–19. Wigge B, Gardeström P. The effects of different ionic conditions on the activity of cytochrome c-oxidase in purified plant mitochondria. In: Moore AL, Beachy RB, editors. Plant mitochondria. New York: Plenum Press; 1987. p. 127–30. Wulff-Zottele C, Gatzke N, Kopka J, Orellana A, Hoefgen R, Fisahn J, Hesse H. Photosynthesis and metabolism interact during acclimation of Arabidopsis thaliana to high irradiance and sulphur depletion. Plant Cell Environ 2010;33:1974–88. Yi H, Galant A, Ravilious GE, Preuss ML, Jez JM. Sensing sulphur conditions: simple to complex protein regulatory mechanisms in plant thiol metabolism. Mol Plant 2010;3:269–79. Yoshimoto N, Inoue E, Watanabe-Takahashi A, Saito K, Takahashi H. Posttranscriptional regulation of high-affinity sulfate transporters in Arabidopsis by sulfur nutrition. Plant Physiol 2007;145(2):378–88. Zhang J, Sun X, Zhang Z, Ni Y, Zhang Q, Liang X, et al. Metabolite profiling of Arabidopsis seedlings in response to exogenous sinalbin and sulfur deficiency. Phytochemistry 2011;72(14-15):1767–78. Zhang S, Chen F, Peng S, Ma W, Korpelainen H, Li C. Comparative physiological, ultrastructural and proteomic analyses reveal sexual differences in the responses of Populus cathayana under drought stress. Proteomics 2010;10(14):2661–77. Zuchi S, Cesco S, Varanini Z, Pinton R, Astolfi S. Sulphur deprivation limits Fedeficiency responses in tomato plants. Planta 2009;230:85–94.