Introduction

Reactive oxygen species (ROS) are generally regarded as harmful products of oxygenic metabolism causing oxidative stress and cell damage. However, in the past years it became apparent that ROS, especially hydrogen peroxide, have a major role in cellular signalling pathways across a wide range of organisms, including plants (Foyer and Noctor 2009). ROS can be generated in plants via the mitochondrial or photosynthetic electron transport chain (ETC), by peroxisomes or can be synthesized in the apoplast by a variety of dedicated enzymes, such as peroxidases and NADPH oxidases (Foyer and Noctor 2009). Once produced, ROS can take part in signalling, although these events are modulated by antioxidants in or around the cell. Some of the key events in plants which are controlled by ROS include stomatal closure (Pei et al. 2000), root growth (Foreman et al. 2003), programmed cell death and the hypersensitive response (Desican et al. 1998).

The plasmalemma of plant cells produces bursts of H2O2 in response to biotic (Vanacker et al. 1998) and abiotic stimuli (Laloi et al. 2004; Piotrovskii et al. 2011). The mechanisms involved in regulation of these bursts are largely unknown but it is speculated that ROS such as H2O2 can also serve as regulators of the redox state of cells allowing controlled oxidation of cell compartments (Foyer and Noctor 2009). In plants the production of ROS initiated by the plasma membrane NADPH oxidase can be used for controlled polymer breakdown leading to wall loosening during extension growth (Schopfer and Liszkay 2006). The redox signals are also integral to the transduction sequences by which changes in the environment modify metabolism and gene transcription (Foyer and Noctor 2009).

ROS perception, signalling and ROS-mediated protein modifications are controlled and modulated by an array of antioxidants. In plants ascorbate (AsA) plays a key role in defence against oxidative stress and is particularly abundant in photosynthetic tissues (Foyer and Noctor 2009). Over 90% of the AsA is localized in the cytoplasm, but a small portion is exported to the apoplast, where it is present in millimolar concentrations. It is by far the most abundant low-molecular-weight antioxidant in the apoplast (Vanacker et al. 1998). Other antioxidants, such as GSH, are either absent from the apoplast or present in low concentrations (Vanacker et al. 1998). In the apoplast AsA is oxidized to monodehydroascorbate (MDHA) by ascorbate oxidase (AO). MDHA is an unstable radical and rapidly disproportionates to yield dehydroascorbate (DHA) and AsA. DHA is then transported into the cytosol through the plasma membrane by a specific carrier that preferentially translocates the oxidized form in exchange for the reduced form, ensuring the continuous flux of reducing power to the cell wall (Horemans et al. 2000). AsA is not only an important antioxidant but also a cofactor of many enzymes (Smirnoff and Wheeler 2000) and a regulator of cell division and growth (Kerk and Feldman 1995). Moreover, AsA is a signal-transducing molecule in plants (Pastori et al. 2003; Foyer and Noctor 2009). Antioxidant defence of the apoplast received lots of attention when the importance of this compartment in cell growth, plant defence, signal transduction and mineral nutrition has become apparent (Foyer and Noctor 2005).

One of the enzymes involved in AsA metabolism, with no clear biological function described to date, is the apoplastic ascorbate oxidase. However, it is believed that AO plays a role in cell elongation because of its extracellular localization and its high activity in rapidly expanding tissues (Kato and Esaka 1999). Several mechanisms whereby AO controls cell growth have been proposed (Smirnoff 2000). MDHA generated from AsA by AO in the apoplast stimulates cell growth through enhanced vacuolization and ion uptake caused by depolarization of the plasma membrane. Moreover, DHA is considered to be responsible for cell enlargement by promoting cell wall loosening (Lin and Varner 1991). AsA is believed to be the most important antioxidant in the apoplast of leaves and stems and that is why its destruction via AO may be important in facilitating cell expansion. Pignocchi and Foyer (2003) showed that expression of AO is regulated by light and by growth-modulating plant hormones. Moreover, they have demonstrated that in transformed tobacco plants enhanced apoplastic AO activity oxidizes the apoplastic AsA pool resulting in stimulation of growth (Pignocchi and Foyer 2003). Taken together, all published data provide the evidence that AsA and AO in the apoplast are key players in both cell division and cell expansion. ROS level, abundance and redox state of the apoplastic ascorbate pool, as well as the enzymes involved in AsA production and turnover play a role in growth control of plants. Thus, plant growth is the outcome of various mechanisms of regulation, signalling and their crosstalk.

The mosaic (MSC16) mutant of cucumber was produced from in vitro cultures by regeneration of the original cucumber B cell line (Malepszy et al. 1996). Crossing the MSC16 mutant with line B showed that the MSC16 phenotype is not associated with either the nuclear or the chloroplast genome (Malepszy et al. 1996). Plants with the MSC16 phenotype have a JLV5 deletion in a non-coding region of the mitochondrial genome, although the precise nature of this mutation remains unknown (Bartoszewski et al. 2004). The MSC16 line has marked phenotypic changes, including a slower growth rate and a leaf mosaic pattern with chlorotic patches (Malepszy et al. 1996). The mitochondrial genome rearrangement in cucumber MSC16 results in a decreased leaf ATP concentration, changes in nucleotides distribution (Szal et al. 2008) and changes in respiratory chain activity, including lower Complex I capacity, increased activities of external NADH dehydrogenases (NDex), higher alternative oxidase (AOX) capacity and protein level (Juszczuk et al. 2007; Juszczuk and Rychter 2009). Cytochemical detection revealed an increased abundance of H2O2 in the mitochondrial membrane and decreased H2O2 content in the apoplast of mesophyll cells in MSC16 leaves (Szal et al. 2009). The mitochondrial mutation also resulted in changes in the antioxidant defence system (Szal et al. 2009). However, these changes in MSC16 metabolism only partially explain its phenotype, especially the altered growth rate.

The present study was undertaken to elucidate the role of the apoplast in formation of the MSC16 phenotype. We showed that a lower content of apoplastic ROS caused by decreased activity of the plasma membrane NADPH oxidase may contribute to altered growth and development of MSC16 cucumber mutant. This can be a result of changes in whole cell energy and redox state caused by impairment of mitochondrial function.

Materials and methods

Plant material and growth conditions

Cucumber seeds (Cucumis sativus L. cv. Borszczagowski) of wild type (WT) or MSC16 mutant were germinated on moist filter paper and 3-day-old seedlings were transferred to boxes with complete Knop medium (Rychter and Mikulska 1990). The nutrient solution was supplemented every day and changed every 4 days. Plants were grown under 16 h photoperiod at 150 μmol m−2 s−1 PAR (daylight and warm white 1:1, LF-40W, Piła, Poland), day/night temperature of 24°C/20°C and 60/70% relative humidity. Leaves from the same developmental stage from about 2-week-old WT and 3-week-old MSC16 plants were taken for analysis.

In vivo detection of H2O2 in plants

H2O2 was visually detected in the leaves of plants by using 3,3-diaminobenzidine (DAB) as substrate. Plants were excised at the base of leaves with a razor blade and supplied with a 1 mg mL−1 solution of DAB, pH 3.8, for 8 h under light at 24°C as described by Orozco-Cárdenas and Ryan (1999).

Cytochemical localization of hydrogen peroxide in leaf mesophyll cells

Cellular H2O2 localization was determined cytochemically from cerium perhydroxide deposition after reaction of CeCl3 with endogenous H2O2 (Bestwick et al. 1997). Positive staining was detected in electron micrographs as the formation of electron-dense deposits. For determination of NADPH oxidase participation in apoplastic ROS production leaf fragments were previously incubated for 1 h with 8 μM DPI. Tissues were then fixed in 2% glutaraldehyde in HEPES buffer (pH 7.3) at room temperature for 2 h. After fixation they were washed three times for 10 min in 0.05 M HEPES buffer at room temperature and post-fixed for 90 min in 1% osmium tetraoxide. After dehydration in an ethanol series (50, 70, 96 and 100%) samples were embedded in Epon 812 resin and polymerized. Sections were made and examined with transmission electron microscope (Leo 912AB). Photographs of the semi-thin sections were taken.

Preparation of PM vesicles

Plasma membrane vesicles were prepared strictly at 4°C (Palmgren 1990). Briefly, whole leaves (50–90 g FW) of MSC16 or WT plants were ground in the presence of 0.6% (w/v) polyvinylpolypyrrolidone (PVPP) with a homogenizing buffer 50 mM Mops-1,3-bis(tris[hydroxymethyl]methylamino) propane (BTP) (pH 7.0) with 330 mM sucrose, 5 mM EDTA, 5 mM dithiothreitol (DTT), 1 mM phenylmethylsulfonyl fluoride (PMSF), 5 mM ascorbate, 0.2% (w/v) bovine serum albumin and 0.2% (w/v) casein. The homogenate was filtered through nylon mesh (240 μm) and centrifuged (10,000×g, 15 min). The supernatant was centrifuged (30,000×g, 55 min), and the resulting precipitate was resuspended with a glass homogenizer in suspension buffer consisting of 5 mM K-phosphate (pH 7.8) with 330 mM sucrose and 2 mM DTT. The homogenate was loaded on a 24 g two-phase system containing 6.7% (w/w) Dextran T500, 6.7% (w/w) polyethylene glycol 4000, 330 mM sucrose, 5 mM K-phosphate (pH 7.8) and 4 mM KCl. After the batch procedure, the resulting upper phase was mixed with a dilution buffer that consisted of 330 mM sucrose, 5 mM Na-phosphate (pH 7.8), and centrifuged (120,000×g, 45 min). This method results in apoplastic-side-out vesicles. To ensure substrate accessibility for NADPH oxidase measurements, the vesicles were turned cytoplasmic-side-out by repeated freezing in liquid nitrogen and thawing in room temperature. In our earlier experiments we found similar effect of both Brij 58 and freezing–thawing technique. The cytoplasmic-side-out PM vesicles were used immediately, or stored otherwise at −80°C until further analysis.

Determination of H+ ATPase activity in PM vesicles

ATPase (EC 3.6.3.6) activity was determined simultaneously with proton pumping (Palmgren and Sommarin 1989). The assay medium consisted of 10 mM Mops-BTP (pH 7.0), 2 mM ATP, 4 mM MgCl2, 140 mM KCl, 1 mM EDTA, 1 mM DTT, 20 μM acridine orange, 0.25 mM NADH, 1 mM PEP, 50 μg mL−1 pyruvate kinase (Roche, 85884429), 25 μg mL−1 lactate dehydrogenase (Sigma, L-2518), and 50 μg mL−1 plasma membrane protein in a total volume of 1 mL. Stock solutions of ATP, EDTA, BSA, and PEP, all pH 7.0 with BTP, were stored in frozen vials and thawed just prior to use. A mixture containing Mops-BTP, ATP, KCl, EDTA, DTT, BSA, and acridine orange was equilibrated at 20°C and a portion of 800 μL was transferred to a disposable 1-mL cuvette, 1 cm light path. Then, 5 μL of membrane suspension was added, followed by 70 μL of a freshly prepared solution of NADH mixed with PEP, 30 μL of a mixture of pyruvate kinase and lactate dehydrogenase. After incubation for 5 min, automatic recording at 15-s intervals at 340 and 495 nm was initiated using a spectrophotometer (Shimadzu UV-160A) that can automatically switch between the two wavelengths. Twenty microliters of 272 mM MgCl2 were added to start the reaction and mixed into the assay solution by stirring for a few seconds with the tip of the pipette. In this assay the rate of ATP hydrolysis was quantified from the rate of NADH oxidation measured at 340 nm after 30 s. Proton uptake into the vesicles was monitored simultaneously as the absorbance decrease at 495 nm of the ΔpH probe acridine orange. ATPase activity was measured in absence and presence of 100 mM Na3VO4, a specific inhibitor of plasma membrane H+ ATPase activity. Proton pumping was determined from the initial slope of acridine orange absorbance quenching.

Determination of cytochrome c oxidase activity

Cytochrome c oxidase (COX, EC 1.9.3.1) activity was measured according to Wigge and Gardeström (1987) in a medium consisting of 0.45 M mannitol, 10 mM phosphate buffer (pH 7.2), 100 mM KCl, 50 μM cytochrome c (reduced with a few crystals of sodium dithionite) and 20 μg plasma membrane protein in the presence of 0.02% (w/v) Triton X-100.

Protein gel blot analyses of H+ ATPase

Protein gel blot analyses were performed using plasma membrane vesicles isolated from MSC16 or WT leaves. Samples of 5, 10 and 20 μg protein per lane, were separated by SDS-PAGE (10% polyacrylamide) according to a standard protocol. The polypeptides were electroblotted onto nitrocellulose membrane and probed with primary antibody overnight at 4°C and 30 min RT. Plasma membrane H+ ATPase (anti-H+ ATPase, Agrisera) antibodies were used as primary antibodies at a dilution of 1/1,000. Anti-rabbit antibodies conjugated to horseradish peroxidase (BIO-RAD) were used as secondary antibodies at a dilution of 1/35,000. Visualization was performed with a chemiluminescent reagent system. Blot was quantified by densitometry using Quantity One 4.6.2 software (Bio-Rad Ltd. USA).

Determination of NADPH oxidase activity

Superoxide anion production by plasma membrane-bound NADPH oxidase (EC 1.6.3.1) was determined using a lucigenin-enhanced chemiluminescence method (Chen et al. 2005). The reaction test tube contained 50 mM phosphate buffer (pH 7.0), 150 mM sucrose, 1 mM EGTA, 5 mM lucigenin and plasma membrane vesicles (about 100 μg of protein). The reaction was started by addition of NADPH to a final concentration of 0.1 mM. Chemiluminometric measurements were performed using Thriatler 425-014 luminometer (Hidex Ltd. Turku, Finland).

Determination of ascorbate oxidase activity

Ascorbate oxidase (EC 1.10.3.3) activity was determined according to Nanasato et al. (2005) with minor modifications. The fourth leaf of WT or MSC16 plants (0.5 g FW) was ground in liquid nitrogen and homogenized with 1 mL of 50 mM K-phosphate buffer (pH 7.0), 1 M KCl, 10 mM 2-mercaptoethanol, 1.5% (w/v) PVPP and 1 mM PMSF. The mixture was allowed to stand on ice for 20 min, vortexed every 3 min and centrifuged at 15,000×g for 20 min at 4°C. The resulting supernatant was used immediately for the enzyme assay. Ascorbate oxidase activity was determined by measuring the decrease in absorbance at 290 nm due to AsA oxidation in the reaction mixture containing 50 mM K-phosphate (pH 5.3), 0.5 mM AsA and 20 μL of leaf extract. Oxidation of AsA was monitored at 25°C (ε = 2.8 mM−1 cm−1).

Extraction of soluble apoplastic components

Soluble apoplastic metabolites were determined in external washing fluid (EWF) by a method similar to that described by Polle et al. (1990). Freshly cut leaves (5 g) were washed three times with distilled water, placed in aluminium foil dishes containing 50 mL of infiltration solution consisting of 50 mM acetate buffer (pH 4.5), 100 mM KCl, and 2 mM CaCl2. To ensure immersion, a second perforated aluminium dish was placed on top of the leaves in the infiltration solution. The infiltration dishes were placed in a vacuum desiccator and the leaves were infiltrated for 5–7 periods of 5 min at about −70 kPa. Then they were blotted gently, loaded into a perforated centrifuge tube (5 mL), and placed in a Falcon tube (14 mL). EWF was recovered by centrifugation (10 min, 2,900×g, 4°C), directly in Falcon tubes containing 100 mL of 0.1 M HClO4 to immediately stop metabolism. Before analysis, sufficient K2CO3 (5 M) was added to each sample of EWF to adjust the pH from 4.0 to 5.0. Between 1.1 and 1.2 mL of EWF was obtained from 5 g of leaves (FW). EWF was used immediately after isolation for ascorbate measurements, in all cases the samples were kept at 4°C until assay. Contamination by cytoplasmic enzymes of the EWF fraction, monitored by the activity of glucose-6-phosphate dehydrogenase (Ros-Barceló et al. 2002) was always less than 0.2%.

Ascorbate determination

Ascorbate was assayed according to the method described by Kampfenkel et al. (1995). In order to measure total ascorbate content, DHA was reduced with DTT. DHA content was calculated from the difference between the concentration of total and reduced ascorbate.

Cell wall preparation

Cell walls from leaves of WT and MSC16 plants were prepared using a method of Wu et al. (1998) modified by Solecka et al. (2008). Fresh leaf tissues were homogenized in 0.05 M HEPES buffer (pH 6.5), containing a mixture of protease inhibitors (PMSF, aprotinin, bestatin, pepstatin A and leupeptine, 1 mM each), filtered through Miracloth and washed several times with cold water. Cell wall proteins were extracted from crude cell wall preparation with HEPES buffer containing 1 M NaCl for 12 h at 4°C (Solecka et al. 2008). Dry matter of crude cell wall preparations after air drying was determined by weighing.

Determination of cell wall guaiacol peroxidase activity

POX (EC 1.11.1.7) activity was analysed in 44 mM acetate buffer (pH 5.0) containing 10 mM guaiacol and 2.2 mM H2O2 at room temperature as described by Van den Berg and Van Huystee (1984). The increase in absorbance at 470 nm was recorded for 3 min after 1 mM H2O2 addition. Enzyme activity was expressed as the average of three replicates in units mg−1 protein−1.

Protein determination

Protein content was measured according to the method of Bradford (1976) with bovine serum albumin (BSA) as a standard.

Statistical analysis

The results presented are the mean values with standard deviations of n (n = 3–11) measurements from 3–5 independent plant cultures. Significance of the results compared to the control was tested using Student’s t test. Asterisk indicates significant differences (P ≤ 0.05).

Results

It was reported earlier (Malepszy et al. 1996; Juszczuk et al. 2007) that MSC16 mutant plants have a considerably slower growth rate and deformed leaves comparing to WT. Cross section of leaves showed that in the mesophyll tissue of MSC16 the area of one cell is often 50–100% greater than of comparable WT leaf cells (Fig. 1). DAB staining of the whole leaves showed less dark brown spots in MSC16 leaves as compared to WT (Fig. 2) indicating lower H2O2 level in MSC16 leaves. Low H2O2 level was found to be mainly located at the cell wall and/or attached to the plasma membrane (Szal et al. 2009). Incubation of leaf tissue with DPI, a potent inhibitor of NADPH oxidase, strongly reduced the production and accumulation of H2O2 in the apoplast of both WT and MSC16 cells (Fig. 3).

Fig. 1
figure 1

Cross-section through leaf blades of WT (a) and MSC16 (b) plants. MSC16 plants have thicker leaf blades with a disturbed structure of palisade and spongy mesophyll. Palisade mesophyll cells of MSC16 plants are bigger, often with irregular shapes (black arrows)

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Fig. 2
figure 2

Localization of H2O2 in leaves of WT (a) and MSC16 (b) plants by DAB staining. Leaves from 2 (WT) or 3 (MSC16) weeks old plants of the same developmental stage were cut and stained with DAB solution

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Fig. 3
figure 3

Cellular localization of H2O2 in mesophyll cells of cucumber MSC16 (a, b) and WT (c, d) leaf blades with CeCl3-staining and transmission electron microscopy before and after DPI treatment. After DPI treatment no H2O2 accumulation was detected in the apoplast of MSC16 (b) and only a little in the apoplast of WT (d); Ch chloroplast, CW cell wall, M mitochondrion, P peroxisome

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NADPH oxidase is located in the plasma membrane, substrate site facing the inner side of plasmalemma (Sagi and Fluhr 2006). To compare the activity of NADPH oxidase we isolated plasma membrane vesicles from leaves of WT and MSC16 plants. The purity of PM vesicles was validated by measuring the activity of enzymes characteristic of other cell compartments. Low activity of the mitochondrial membrane marker cytochrome c oxidase was detected (about 30 nmol min−1 mg−1 protein). The contamination with tonoplast was checked as ATPase activity, measured in the presence and absence of an inhibitor. Vanadate, which is a specific inhibitor of PM ATPase activity (Palmgren and Sommarin 1989) inhibited enzyme activity by more than 60%. To check if the plasma membrane vesicles isolated from both MSC16 and WT leaves had comparable protein content, H+ ATPase protein level was measured. Arabidopsis PM H+ ATPase antibody detected a single band at approximately 95 kDa in PM vesicle proteins, showing a comparable protein amount in MSC16 and WT PM vesicles as estimated by densitometric measurements (Fig. 4). In PM vesicles isolated from MSC16 leaves NADPH oxidase activity was about 30% lower compared to PM vesicles isolated from WT leaves (Fig. 4).

Fig. 4
figure 4

NADPH oxidase activity in the plasma membrane vesicles isolated from WT and MSC16 leaves, measured using a lucigenin-enhanced chemiluminescence method. Each value represents the mean ± SD of five independent measurements. In the upper right corner protein determination of plasma membrane H+ ATPase from PM vesicles was given as a control. Eight micrograms of protein was loaded per lane, separated by SDS-PAGE, immunoblotted and visualized by chemiluminescence

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Cell wall peroxidases are also responsible for apoplastic ROS production (Wojtaszek 1997; Ros-Barceló 1998) and, similarly to the plasma membrane NADPH oxidase, their activity could also be inhibited by DPI. To determine if peroxidase activity may contribute to the decreased H2O2 content we have isolated cell walls from WT and MSC16 plants and found that guaiacol peroxidase activity was low and comparable in both plants (0.7–0.8 U min−1 mg−1 protein).

AsA is an important component of the complex network of control and interactions that regulates growth. Therefore, the abundance and redox state of the major soluble antioxidant-ascorbate was estimated in soluble apoplastic components isolated from WT and MSC16 leaves and compared to the values found in leaf homogenates. Apoplastic ascorbate (AsA + DHA) accounts only for 2–3% of total ascorbate measured in homogenates in MSC16 or WT leaves (Szal et al. 2009). Total ascorbate concentration was about 14% higher in MSC16 EWF extracts and the reduced form (AsA) was substantially more abundant than in WT EWF extracts (Fig. 5).

Fig. 5
figure 5

Ascorbate concentration in leaf soluble apoplastic components from WT and MSC16 plants. Total (AsA + DHA) and reduced (AsA) ascorbate were determined separately using a colorimetric method. Values are means ± SD from three replicates. Marked significance relates to the level of both AsA and DHA

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Ascorbate oxidase (AO), the enzyme responsible for ascorbate metabolism in the apoplast is present solely in the apoplast; therefore, its activity was detected in whole-leaf extracts. The AO activity measured as the oxidation of AsA to DHA was about 40% higher in MSC16 leaf extracts compared to WT (Fig. 6).

Fig. 6
figure 6

Ascorbate oxidase activity in leaf samples of WT and MSC16 plants. Each value represents the mean ± SD of three independent measurements

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Discussion

In addition to earlier described effects of mitochondrial mutation on MSC16 leaf carbon and energy metabolism which may affect growth (Juszczuk et al. 2007; Szal et al. 2009, 2010), our data indicate also the possible involvement of apoplastic ROS. ROS level in the apoplast is the result of the activities of plasma membrane-located NADPH oxidases (Low and Merida 1996; Lamb and Dixon 1997) and cell wall-located peroxidases (Wojtaszek 1997). Diphenyleneiodonium (DPI) is known to be a potent inhibitor of the NADPH oxidase and other flavin-containing oxidases and have been used extensively to inhibit production of ROS in plant systems (Bolwell et al. 1995). Except plasma membrane NADPH oxidase, DPI inhibits cell wall peroxidases (Ros-Barceló 1998) and respiratory chain Complex I (Fato et al. 2009). Differences in apoplastic ROS concentration in WT and MSC16 seem not to be dependent on mitochondrial ROS production since ROS production by isolated mitochondria was similar in both plants (Szal et al. 2009) also cell wall peroxidase activity was not different in WT and MSC16. Although the involvement of chloroplast and peroxisomes in ROS production cannot be ignored our previous data indicated that mitochondrial mutation in MSC16 does not significantly affect photosynthesis and photorespiration (Juszczuk et al. 2007). All together it seems that the lower H2O2 level detected in the apoplast of MSC16 cucumber leaves may result from the decreased NADPH oxidase activity.

In the apoplast the plasma membrane NADPH oxidase is using cytosolic NADPH as an electron donor and generates O ·−2 and subsequently H2O2 by the activity of SOD. We did not estimate apoplastic SOD activity but the activities of Cu/ZnSOD isoforms 1, 2 and 3 were higher in MSC16 extracts while the activity of MnSOD was similar in both WT and MSC16 (Szal et al. 2009). NADPH oxidase-derived ROS are required during the growth of organs, for example during leaf expansion (Rodriguez et al. 2002). The increase in NADPH oxidase activity observed in vivo during cold treatment (Piotrovskii et al. 2011) or pathogenesis response (Neill et al. 2002) may be due to the changes in substrate (NADPH) availability or modulation by cytoplasmic factors. The mitochondrial genome rearrangement in cucumber MSC16 results in changes in respiratory chain activity, including increased activities of external NAD(P)H dehydrogenases (NDex) (Juszczuk et al. 2007). Moreover, in MSC16 plants cytosolic NADP(H) pool is lower and more oxidized than in WT (Szal et al. 2008), which in vivo may have influence on NADPH oxidase activity in MSC16. As all the measurements were performed in vitro with saturating levels of substrate, we can expect that NADPH oxidase activity in vivo may be even lower since it is dependent not only on substrate availability but also on enzyme protein level or other regulating factors like Rho-related GTPases (Baxter-Burrell et al. 2002). Therefore, we suggest that the lower NADPH oxidase activity in cucumber mutants, resulting in low apoplastic ROS production, may influence MSC16 plant growth.

The total pool of foliar ascorbate (reduced plus oxidized) was about 35% higher in MSC16 leaves than in WT leaves and the greater total abundance of ascorbate in MSC16 leaves was accompanied by a large increase in the oxidized form dehydroascorbate (DHA) (Szal et al. 2009). Cell division is sensitive to changes in apoplastic AsA, MDHA and DHA. AsA concentrations present in the cucumber leaf apoplast, which ranged in WT and MSC16 from 18 to 22 nM, respectively (Fig. 5) are comparable to the AsA levels in the apoplast of barley (16 nM) and oat leaves (10 nM) (Vanacker et al. 1998). Total ascorbate concentration was about 14% higher in MSC16 EWF extracts and the reduced form (AsA) was substantially more abundant than in WT (Fig. 5). The apoplastic ascorbate pool of MSC16 plants was in 94% reduced, suggesting a very efficient system of DHA translocation from the apoplast to the cytosol by a carrier-mediated AsA/DHA transport system (Horemans et al. 2000). This might explain, at least in part, why total foliar ascorbate in MSC16 plants was almost in 50% oxidized (Szal et al. 2009).

In the apoplast of MSC16 plants AO activity was about 40% higher than in WT plants (Fig. 6). An increased AO activity leads to enhanced cell expansion in transgenic tobacco plants expressing AO in sense orientation (PAO) (Pignocchi et al. 2003). The apoplast of PAO plants also contained higher amounts of ascorbate (AsA + DHA) than the apoplast of untransformed plants (Pignocchi et al. 2003). No DHA accumulation was observed despite about 40% higher AO activity in MSC16 plants than in the apoplast of WT plants. The positive correlations between AO activities, apoplastic DHA and plant growth does not take place in MSC16 plants. It is worth saying, that oxidation of apoplastic AsA alone favours enhanced growth in transgenic tobacco overexpressing AO but does not affect cell division (Pignocchi and Foyer 2003). They also speculate that enhanced DHA accumulation could signal metabolic arrest and inhibit cell division, pointing out at least two possibilities. One of them is that DHA accumulation alone is insufficient to arrest mitosis and another possibility is that a second signal must be involved (Pignocchi and Foyer 2003). In this case it is possible to attribute the AO-dependent enhancement of growth to cell elongation alone. This is exactly what takes place in MSC16 plants. The observed greater dimensions of mutant’s mesophyll cells (Fig. 1) as compared with WT might be a result of enhanced cell expansion due to unsettled stiffening of the cell wall. The cell wall plays an important role in cell expansion; loosening allows cells to grow, whereas wall cross-linking can inhibit this process. There is evidence that ROS are involved in both processes (Ros-Barceló et al. 2002; Liszkay et al. 2004).

Mitochondrial genome rearrangement resulting in altered activity of ETC complexes, implies a radical alteration in whole cell redox metabolism; e.g. adenylate concentration, pyridine nucleotide redox state (Szal et al. 2008) and ROS/antioxidants homeostasis (Szal et al. 2009). Changes in the energy state of leaf cells can influence the activity of a number of specific proteins that regulate growth processes, such as NADPH oxidase and AO or other. In addition to the earlier described effects of mitochondrial mutation on MSC16 leaf carbon and energy metabolism which may affect growth (Juszczuk et al. 2007; Szal et al. 2009, 2010) our data indicate the possible involvement of apoplastic ROS in these processes.