Journal of Experimental Botany

Halford, Nigel G.

doi: 10.1093/jxb/erp311pmid: 19854798

Those of us working and living in temperate countries often disregard the effects of heat stress on crop yield and quality, possibly to the frustration of our colleagues from hotter parts of the world. That is changing as global warming is predicted to increase the frequency and severity of ‘heat-waves’ in temperate zones (Semenov, 2007; Semenov and Halford, 2009) but the paper from Ginzberg and colleagues published in this issue (Ginzberg et al., 2009) reminds us that heat stress is already a problem of economic significance. The paper describes the effects of heat stress on potato tuber skin quality: a soil temperature of 33 °C causes russeting in which the skin thickens, then cracks, resulting in a rough skin texture and a reduced value of the crop. It is another example of how the application of modern techniques such as transcriptomics can reveal that something like the roughening of the skin of potato tubers may be the visible indicator of quite dramatic changes in gene expression occurring within a plant as it responds to a stress. The results of the study add to the growing evidence that stress signalling pathways are not independent, but interact to form networks. Hence, many of the genes that showed changes in expression in this study in response to heat stress have previously been shown to be involved in other stress responses. The concept of interacting signalling networks has been discussed previously with respect to, for example, jasmonate signalling (Lorenzo and Solano, 2005), as well as signalling pathways involving mitogen-activated protein kinases (MAPKs) (Colcombet and Hirt, 2008) and sucrose nonfermenting-1 (SNF1)-related protein kinases (Halford and Hey, 2009). Indeed, the ‘network’ model is replacing the ‘pathway/cascade’ model as the best fit for eukaryotic signalling systems (Halford and Hey, 2009). The study highlights one of the advantages of using no-preconception, ‘omic’ techniques in that sometimes unexpected interactions are revealed. For example, a number of genes previously shown to be involved in the development of symbiotic or protective membranes during plant/microbe interactions were found to increase in expression in response to heat stress. Furthermore, the expression of many genes encoding transcription factors associated with the regulation of cell proliferation, orientation, and differentiation was affected. It is doubtful if these genes would have been selected for analysis if a candidate-gene approach had been taken. On the other hand, the study also draws attention to the problem of turning all of the information that is produced in an ‘omics’-based project into knowledge: investigating all of the genes that showed a change in expression in response to the stress would require staff and resource levels beyond those available to most research teams. So, as with almost all ‘omics’-based studies, a subset of genes was chosen for further study, the researchers choosing to focus on genes encoding transcription factors. This is not surprising given the potential for transcription factors to have wide-ranging effects on plant physiology, but is still somewhat arbitrary. Such decisions are unavoidable and familiar to many of us, but to some extent defeat the object of using a no-preconception approach. A failing of a number of studies on the effects of heat stress has been to assume that heat and drought stress are synonymous. In temperate countries, of course, periods of hot weather are often also dry, while the effects of drought in winter, when it is cold, are minimized by the fact that plants are dormant or semi-dormant. However, in many parts of the world, plants have to cope with hot, wet conditions, and where plants do have to endure periods of dry weather they may not be affected even by severe drought if they are able to control water loss effectively and access water sources from deep in the soil. Such plants may be affected by abnormally high temperatures much more quickly than they are affected by drought (reviewed in more detail by Semenov and Halford, 2009). It is notable that Ginzberg and colleagues report that three transcription factors associated with drought responses were actually down-regulated in heat-stressed tubers. It is important that this is taken on board so that genuine heat stress tolerance markers can be identified. Otherwise, new genotypes may be developed to cope with high temperatures and fail because the wrong markers have been selected for. Finally, Ginzberg and colleagues resisted invoking the issue of climate change in their paper so I will do it for them. Heat stress has been shown to reduce quality and yield of other major crops, affecting, for example, fertility and seed development of wheat (Gooding et al., 2003), maize (Wilhelm et al., 1999), and rice (Jagadish et al., 2007). At the same time, periods of destructively high temperature, which have occurred in the past perhaps once every century, are predicted to become much more frequent by the end of this century, occurring perhaps once or twice per decade (Semenov, 2007; Semenov and Halford, 2009). Clearly, therefore, understanding how plants respond to heat stress and how heat tolerance can be improved is of the highest importance. Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council (BBSRC) of the United Kingdom. References Colcombet J,  Hirt H.  Arabidopsis MAPKs: a complex signalling network involved in multiple biological processes,  Biochemical Journal ,  2008, vol.  413 (pg.  217- 226) Google Scholar CrossRef Search ADS PubMed  Ginzberg I,  Barel G,  Ophir R,  Tzin E,  Tanami Z,  Muddarangapa T,  de Jong W,  Fogelman E.  Transcriptomic profiling of heat-stress response in potato periderm,  Journal of Experimental Botany ,  2009, vol.  60 (pg.  4411- 4421) Google Scholar CrossRef Search ADS PubMed  Gooding MJ,  Ellis RH,  Shewry PR,  Schofield JD.  Effects of restricted water availability and increased temperature on the grain filling, drying and quality of winter wheat,  Journal of Cereal Science ,  2003, vol.  37 (pg.  295- 309) Google Scholar CrossRef Search ADS   Halford NG,  Hey SJ.  SNF1-related protein kinases (SnRKs) act within an intricate network that links metabolic and stress signalling in plants,  Biochemical Journal ,  2009, vol.  419 (pg.  247- 259) Google Scholar CrossRef Search ADS PubMed  Jagadish SVK,  Craufurd PQ,  Wheeler TR.  High temperature stress and spikelet fertility in rice (Oryza sativa L.),  Journal of Experimental Botany ,  2007, vol.  58 (pg.  1627- 1635) Google Scholar CrossRef Search ADS PubMed  Lorenzo O,  Solano R.  Molecular players regulating the jasmonate signalling network,  Current Opinion in Plant Biology ,  2005, vol.  8 (pg.  532- 540) Google Scholar CrossRef Search ADS PubMed  Wilhelm EP,  Mullen RE,  Keeling PL,  Singletary GW.  Heat stress during grain filling in maize: effects on kernel growth and metabolism,  Crop Science ,  1999, vol.  39 (pg.  1733- 1741) Google Scholar CrossRef Search ADS   Semenov MA.  Development of high resolution UKCIP02-based climate change scenarios in the UK,  Agricultural and Forest Meteorology ,  2007, vol.  144 (pg.  127- 138) Google Scholar CrossRef Search ADS   Semenov MA,  Halford NG.  Identifying target traits and molecular mechanisms for wheat breeding under a changing climate,  Journal of Experimental Botany ,  2009, vol.  60 (pg.  2791- 2804) Google Scholar CrossRef Search ADS PubMed  © The Author [2009]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved. For Permissions, please e-mail: [email protected]

Kandeler, Riklef;Ullrich, Wolfram R.

doi: 10.1093/jxb/erp253pmid: 19752049

DECEMBER: Cross-roses and rose crosses December is a quiet and dark month for plants in the Northern temperate zone and so we turn to some ornamental and symbolic forms of the rose flower. Of all flowers, the rose has been adapted most widely and universally to contribute to symbolism and decoration. We choose some fine examples from early Christian architecture. The motif of incorporating a rose into the sign of the cross, or of adding a cross to the image of a rose, has been used widely in the later Mediterranean antiquity. However, its use as a symbolic decoration reached a peak in some of the early churches of the Byzantine Empire, which embraced most lands around the Mediterranean Sea in the 5th and 6th centuries AD. The cross-rose motif can be seen to take several forms: cross-roses with rose flowers in a frontal view divided symmetrically into four parts by a cross (Fig. 1, left), and as four combined young rose flowers in the angles of a cross, usually presented in lateral view (Fig. 1, middle and right). Before they became adopted by the Christian church they were used in ornamental floor mosaics, and on walls and ceilings throughout the late Roman Empire. Examples can be found in Antioch, Cyprus, Tunisia, and Algeria. In the early centuries of the Christian church the cross symbol was avoided because it was associated with crucifixion and death (Murray and Murray, 2004). However, as noted in an earlier article of this series (October), the rose was to become associated with the Virgin Mary and by the 5th century AD cross-roses and rose crosses had been adopted by the Christians. Fig. 1. View largeDownload slide Simplified forms of a cross-rose (left), a stylized form of a single tetramerous rose flower with a cross; and two types of rose crosses: middle: more similar to a cross-rose; right: the composition of four young, half-opened rose flowers in side view is more pronounced (Ullrich). See also Fig. 2. Fig. 1. View largeDownload slide Simplified forms of a cross-rose (left), a stylized form of a single tetramerous rose flower with a cross; and two types of rose crosses: middle: more similar to a cross-rose; right: the composition of four young, half-opened rose flowers in side view is more pronounced (Ullrich). See also Fig. 2. The incorporation of the rose flower, as a sign of the birth of Christ, into a symbol with the cross, a sign of His death, mixed with the beauty and symmetry of each may explain the prevalence of this motif. Other ideas have been put forward, such as those of Kandeler (2006) who considered that cross-roses were an apotropaion (protection symbol). An old, pre-Christian example of a cross-rose can be seen in a floor mosaic from Pergamum, originating from the 2nd century BC (Fig. 2, October of this series). In the middle of a garland with naturalistic double roses, rose buds and foliage, there is a cross-rose with a dark centre – almost like a foreign element. This conspicuous sign presumably was to complement the fullness of life of the garland with an apotropaion, i.e. with a protective function. Fig. 2. View largeDownload slide Mosaic floor with three-pointed forked cross-roses (round fields) and various forms of rose crosses (interspaces). St Pantaleon, Aphrodisias, Cilicia, 4th century AD (Budde, 1987). Fig. 2. View largeDownload slide Mosaic floor with three-pointed forked cross-roses (round fields) and various forms of rose crosses (interspaces). St Pantaleon, Aphrodisias, Cilicia, 4th century AD (Budde, 1987). Cross-roses were particularly appreciated as symbolic ornaments by the Byzantine imperial court. When, in the 5th century, St Mary's church in Thessaloniki was constructed, certain arcade arches were decorated with mosaics of cross-roses together with lotus flowers, lilies, and ears of grain. In other cases a combination of cross-roses with rose crosses created a rhythmic pictorial, yet symbolic design feature in the floor mosaics as found at St Pantaleon in Cilicia (Turkey; Fig. 2) (Budde, 1987). Such designs incorporating cross-roses are elaborated further in Ravenna, as at the tomb of Galla Placidia (5th century). In San Vitale, San Apollinare Nuovo, and in San Apollinare in Classe (Ravenna), mosaics from the middle of the 6th century AD show figures with a background characterized by roses and lilies. In San Vitale, apart from the figure mosaics that cover all the walls of the interior, cross-roses appear in many ornamental bands and in the arches in the choir of the church combined with symmetric pairs of horns of plenty (Fig. 3). These cross-roses show a rather complex design, dark crosses and forks at the end of the arms (forked crosses). Fig. 3. View largeDownload slide Cross-roses with forked crosses combined with horns of plenty. S. Vitale, Ravenna, mosaic on a band-shaped arch, early 6th century AD (Kandeler). Fig. 3. View largeDownload slide Cross-roses with forked crosses combined with horns of plenty. S. Vitale, Ravenna, mosaic on a band-shaped arch, early 6th century AD (Kandeler). Whether used as an ornament or as a symbol it is clear that the rose and cross were images used creatively, sometimes reduced to minimalist symmetry, other times elaborated. It is interesting to reflect on how well these motifs still work as metaphors in the diversity of modern life. References Budde L. ,  St Pantaleon von Aphrodisias in Kilikien ,  1987 Recklinghausen Aurel Bongers Kandeler R.  Symbolism of plants and colours,  Abhandlungen der Zoologisch-Botanischen Gesellschaft in Österreich ,  2006, vol.  Vol. 33A  Vienna Murray P,  Murray L. ,  Oxford dictionary of Christian art ,  2004 Oxford University Press © The Author [2009]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved. For Permissions, please e-mail: [email protected]

Kandeler, Riklef;Ullrich, Wolfram R.

doi: 10.1093/jxb/erp266pmid: 19755571

EPILOGUE We close the year by looking over our series and by reflecting, with some observations, on creativity, reductionism, and diversity. In the contributions on plant symbolism of this past year only a few examples could be given. They show that, over the millennia, many more relationships between humans and plants have existed than just their use for food, technical purposes, and decoration. It applies to all the plants discussed in these 12 short articles that they were regarded as helpers for the conservation, strengthening, and renewal of life. Their magic forces were believed to offer help for the individual, to increase vitality, and to enable the transition to eternal life. In many cases, medicinal application and the belief in metaphysical forces were closely related to each other. Formal similarity played a role in the belief in healing effects, as for example, with Hepatica triloba and the human liver or the lichen Lobaria pulmonaria with the structure of the lungs. Trust in metaphysical forces led to the belief that symbolism had protective (apotropaic) power. Since the 16th century, but mainly in the time of Enlightenment in the 17th and 18th centuries, the scientific background of botany and of the enormous diversity of the plant kingdom markedly increased and with this the concept of medicine. The arrival of modern medicine and pharmacology finally led to a loss of the transcendental background of symbolism within the 20th century. During the period of Romanticism in the early 19th century some of the mystic ideas were captured into elements of culture which continue to play a role in spiritualistic and esoteric movements to this day. The development of the biological sciences in the 20th century went so far in the direction of reductionism that the biologists’ sense of diversity became suppressed. Diversity and the equilibrium between all living organisms worldwide are again regarded as being very important for biology. Pharmaceutical research is again looking for natural substances in endangered species and habitats. Nevertheless, most of us still know the symbolic implications behind some of our beautiful flowers when we communicate with each other: red roses, yellow flowers, white lilies etc., a symbolism that, of course, will vary with people's cultural background. In a last and, to some extent, summarizing picture of this series, a pretty combination of many symbolic flowers is shown in the ‘Garden of Paradise’ (hortus conclusus) painted by an unknown artist in about 1410 (Fig. 1). The Christian context is obvious: Mary with a book is sitting in the upper centre, in front of her the Christ child is playing a psaltery, held by St Catherine, while St Dorothy is plucking cherries in the upper left and St Barbara is ladling water from a basin. At the right-hand side, Archangel Michael is sitting behind St George, who is characterized by a small dragon lying at the lower edge of the painting, and St Oswald is gripping the trunk of the tree between them. As a possible interpretation of the painting, some authors consider it as showing the reception of a knight with his page in paradise, guided by the Archangel Michael who is the usual guide of souls (Vetter, 1965; Wolters, 1932). The list of symbolic plants in the painting is long, most of them shown growing around the saints. Among them are: columbine (Aquilegia), speedwell (Veronica), lady's mantle (Alchemilla), daisy (Bellis), wallflower (Cheiranthus), periwinkle (Vinca), clover (Trifolium), white lily (Lilium), spring snowflake (Leucojum), lily-of-the-valley (Convallaria), strawberry (Fragaria), mallow (Malva), tansy (Chrysanthemum), carnation (Dianthus), peony (Paeonia), rose (Rosa), cowslip (Primula), iris (Iris), mustard (Sinapis), dead-nettle (Lamium), violet (Viola), and plantain (Plantago). For us in the 21st century most of them may appear as decoration rather than as the hidden language of flowers of earlier centuries. Fig. 1. View largeDownload slide ‘Little Garden of Paradise’ with Mary and the Child in the centre and various saints and many symbolic flowers surrounding them (Master of the Garden of Paradise, 1400–1420 AD, Städel Museum, Frankfurt/Main, loan from Historisches Museum). For details see text. Fig. 1. View largeDownload slide ‘Little Garden of Paradise’ with Mary and the Child in the centre and various saints and many symbolic flowers surrounding them (Master of the Garden of Paradise, 1400–1420 AD, Städel Museum, Frankfurt/Main, loan from Historisches Museum). For details see text. References Vetter EM.  Das Frankfurter Paradiesgärtlein,  Heidelberger Jahrbücher IX ,  1965(pg.  102- 146) Wolters A. ,  1932  Oberrheinische Meister von Anfang des 15. Jahrhunderts. Das Paradiesgärtlein. In: Meisterwerke alter Malerei im Städelschen Kunstinstitut, Frankfurt © The Author [2009]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved. For Permissions, please e-mail: [email protected]

Chaki, Mounira; Valderrama, Raquel; Fernández-Ocaña, Ana M.; Carreras, Alfonso; López-Jaramillo, Javier; Luque, Francisco; Palma, José M.; Pedrajas, José R.; Begara-Morales, Juan C.; Sánchez-Calvo, Beatriz; Gómez-Rodríguez, María V.; Corpas, Francisco J.; Barroso, Juan B.

doi: 10.1093/jxb/erp263pmid: 19717529

Gandin, Anthony; Lapointe, Line; Dizengremel, Pierre

doi: 10.1093/jxb/erp255pmid: 19710178

Abstract Mechanisms that allow plants to cope with a recurrent surplus of carbon in conditions of imbalance between source and sink activity has not received much attention. The response of sink growth and metabolism to the modulation of source activity was investigated using elevated CO2 and elevated O3 growth conditions in Erythronium americanum. Sink activity was monitored via slice and mitochondrial respiratory rates, sucrose hydrolysis activity, carbohydrates, and biomass accumulation throughout the growth season, while source activity was monitored via gas exchanges, rubisco and phosphoenolpyruvate carboxylase activities, carbohydrates, and respiratory rates. Elevated CO2 increased the net photosynthetic rate by increasing substrate availability for rubisco. Elevated O3 decreased the net photosynthetic rate mainly through a reduction in rubisco activity. Despite this modulation of the source activity, neither plant growth nor starch accumulation were affected by the treatments. Sucrose synthase activity was higher in the sink under elevated CO2 and lower under elevated O3, thereby modulating the pool of glycolytic intermediates. The alternative respiratory pathway was similarly modulated in the sink, as seen with both the activity and capacity of the pathway, as well as with the alternative oxidase abundance. In this sink-limited species, the alternative respiratory pathway appears to balance carbon availability with sink capacity, thereby avoiding early feedback-inhibition of photosynthesis in conditions of excess carbon availability. Alternative pathway, bulbous plant, carbohydrate, carbon allocation, carbon metabolism, Erythronium americanum, respiration, sink limitation Introduction During growth, source tissues are photosynthetically active and export synthesized carbohydrates to photosynthetically less active or inactive sink tissues, such as roots, fruits, tubers or bulbs (Dickson, 1991). Sinks are thus characterized by a net importation of carbohydrates, which are used for growth, maintenance, and storage. The ability of a sink organ to import assimilates, also referred to as sink strength, is determined by both sink size and sink activity (Farrar, 1993). Strengths of the different sinks define, in turn, carbon (C) allocation patterns at the whole plant level. C allocation patterns change throughout the season, following plant developmental stages, but many abiotic factors can also modify C allocation pattern through changes in the amount of C fixed by the plant. C allocation in plants with storage organs has often been studied through modulation of the source activity by altering either light levels (Kehr et al., 1998) or gas concentrations (i.e. increasing level of CO2 and O3, Andersen, 2003; Balaguer et al., 1995). Stimulation of source activity translates into higher photosynthetic rates, leading to an increased supply of carbohydrates. Excess C is usually allocated to the sinks, leading to more accumulation of reserves in the storage organs (Balaguer et al., 1995). This is the case for source-limited plants. In sink-limited species, photosynthetic rates are modulated more extensively by sink C-demand than by abiotic factors (Sawada et al., 2003), leading to complex physiological and biochemical controls of source activity. A decrease of assimilate utilization by sinks generally leads to the accumulation of sucrose or starch in leaves, which then decreases photosynthesis through feedback inhibition (Paul and Foyer, 2001). However, sugar accumulation in either source or sink organs can also stimulate respiration (Amthor, 1991) and avoid feedback inhibition of different C metabolism pathways. Indeed, as respiration is central to all C metabolism pathways, it could play a key role in C exchanges between source and sink. Increasing respiratory rates can cause an over-reduction of some of the electron transport chain components (Turrens, 2003). In the terminal step of the respiratory process, electrons can either pass along the phosphorylating cytochrome pathway or the non-phosphorylating alternative oxidase pathway. It has been suggested that the alternative pathway has the ability to use excess ubiquinone electron pools, thereby avoiding over-reduction of the electron transport chain, which could lead to the synthesis of reactive oxygen species (ROS) (Moller, 2001; Rich and Bonner, 1978). This homeostatic regulation is linked to the hypothesis that the alternative pathway acts as an ‘energy overflow’ conduit for the cytochrome pathway (Lambers, 1982). The main factors that determine electron partitioning between the cytochrome and alternative pathways are the ratio of reduced ubiquinone to total ubiquinone pools (Wagner et al., 1998), the amount and redox state of alternative oxidase (AOX) proteins (Umbach and Siedow, 1993), the presence of α-keto acids (Millar et al., 1993; Umbach et al., 1994), and the availability of ADP and Pi (Juszczuk et al., 2001). The levels of these specific metabolites can vary with developmental stage and environmental conditions. However, modulation of the different respiratory pathways has only been well-documented for source organs (Gonzàlez-Meler et al., 2001; Millenaar and Lambers, 2003). Information is scarce for sink organs, especially in relation to source activity. The present study attempted to modulate the source activity in Erythronium americanum Ker Gawl. (trout lily) and to study its impact on both source and sink C metabolism. Source activity was modulated using different environmental gas conditions: elevated CO2 to increase C fixation and elevated O3 to reduce rubisco activity. Due to its simple morphology, E. americanum is very close to a theoretical source–sink model and, thus, is an interesting biological model in which to study whole-plant C allocation. Indeed, 90% of individuals in this species have only one leaf and only one bulb (Blodgett, 1910), corresponding to one source and one strong sink. Erythronium americanum is an abundant spring geophyte of North American maple forests, whose epigeous development begins early in the spring and takes place over a short period, from snow melt to canopy closure (Muller, 1978; Taylor and Pearcy, 1976). Carbohydrate storage in the bulb is renewed during this period and when completed, appears to induce leaf senescence (Lapointe, 2001). Thus, plant growth becomes rapidly sink-limited as the bulb reaches its final size. Source activity was monitored via gas exchanges, rubisco and phosphoenolpyruvate carboxylase (PEPc) activities, respiratory rates, and sugar concentrations, whereas sink activity was monitored via respiratory rates, sucrose hydrolysis activity, carbohydrate concentrations, and plant biomass. This study should help unravel some of the regulatory mechanisms that allow sink-limited species to cope with changes in C availability. Materials and methods Plant material and growing conditions Bulbs of E. americanum were collected in September in a maple forest near Saint-Augustin-de-Desmaures (QC, Canada; 46° 48′ N, 71° 23′ W). Bulbs of similar biomass (0.35–0.40 g fresh weight) were selected and planted in plastic pots containing Turface (calcined clay granules, Applied Industrial Materials Corp., Buffalo Grove, IL, USA) as substrate. Plants were kept in a cold chamber for 5 months of cold stratification at 4 °C and then randomly transferred into eight phytotron chambers. Four different gas treatments were applied to these chambers: control, with 390 mmol mol−1 of CO2 and 0 nmol mol−1 of O3 (charcoal-filtered air); elevated CO2, with 1000 mmol mol−1 of CO2; elevated O3, with 80 nmol mol−1 of O3; and elevated CO2+O3, with 1000 mmol mol−1 of CO2 and 80 nmol mol−1 of O3. Other growth parameters were held constant: photoperiod, 14 h; air temperature, 18/14 °C day/night; relative humidity, 75%; and photon flux density (PPFD), 350 μmol m−2 s−1. Ambient air in each chamber was analysed continuously by an ozone analyser (O341M, Environment SA, Paris, France) and a CO2 analyser (WMA2 PPsystems, Stotfold, UK). Plants were watered daily and fertilized weekly with 10% Hoagland's solution to ensure optimal growth (Lapointe and Lerat, 2006). Gas exchange measurements Net CO2 photosynthetic rate (Pn) was measured under phytotron conditions (CO2, temperature, and RH) using a Li−Cor 6400 Portable Photosynthesis System (Li−Cor Inc, Lincoln, NE, USA). Light conditions during the measurements were constant at 350 μmol m−2 s−1, with air flow at 200 μmol s−1. Measurements were randomized between treatments from 10 h to 13 h. Pn was measured on six plants per treatment (three per chamber) at leaf unfolding (t1, day 5), at the initiation of leaf senescence (t8, day 21), at complete leaf senescence (t9, day 28), and every 2 d from t1 to t8. Plant growth measurements Three plants per chamber were harvested on the same days that Pn was measured, plus one extra harvest at the time plants were moved to the chamber (t0). Leaves, bulbs, and roots were dried for 24 h at 70 °C and weighed separately. Enzyme extraction and assays Measurements were done on four plants per chamber that were harvested 4, 8, 12, 16, and 20 d after transfer to the growth chamber. Leaves and bulbs from each plant were separated and immediately frozen in liquid nitrogen. Tissues were then stored at –80 °C until extraction. Frozen leaf and bulb tissues (300 mg FW) were ground in liquid nitrogen with a mortar and pestle. Crude enzyme extractions were performed at 4 °C according to Fontaine et al. (2002). Enzymatic activities were determined spectrophotometrically (ELX 808 Microplate reader, Bio-Tek instruments, St-Quentin-en-Yvelines, France) in coupled reactions by monitoring NADH oxidation at 340 nm. Measurements of rubisco and PEPc activities were carried out according to Fontaine et al. (2002). Cytoplasmic invertase was assayed in 200 μl of 50 mM HEPES–NaOH (pH 7) containing 2.8 mM ATP, 1.2 mM NADP, 12 U ml−1 hexokinase (EC 2.7.1.1), 3.5 U ml−1 phosphoglucoisomerase (EC 5.3.1.9), 1.75 U ml−1 glucose-6-phosphate dehydrogenase (EC 1.1.1.49), and 10 μl of enzymatic extract. The reaction was initiated by adding 100 mM sucrose. A similar medium with 2 mM UDP was used for the Susy assay. Susy activity was estimated by difference between the reactions with and without UDP. Controls without substrate addition were run with all assays. Carbohydrate concentrations The leaf and bulb of three plants per chamber were analysed for starch, sucrose, and glucose-fructose concentrations at 0, 4, 8, 12, 20, and 24 d after transfer to the growth chambers. Carbohydrates were estimated according to Blakeney and Mutton (1980). Frozen tissues were lyophilized for 24 h and weighed before maceration in a solution of methanol, chloroform, and water (12:5:3 by vol.) for 20 min at 65 °C. The mixture was ground with a Polytron (Kinematica, Lucerne, Switzerland) and centrifuged at 3500 rpm for 10 min at 4 °C. Starch contained in the pellet was gelatinized in boiling water for 90 min and then hydrolysed at 55 °C for 60 min in the presence of amyloglucosidase. The supernatant was analysed before and after invertase digestion to estimate reducing sugars and sucrose concentrations, respectively. Finally, all reducing sugars were quantified colorimetrically at 415 nm after reaction with p-hydroxybenzoic acid hydrazide (Sigma Chemical Co., St Louis, MO, USA). Slice respiration measurements Respiration was recorded on leaves and bulbs of six plants per treatment at 4, 8, 12, 16, and 20 d following transfer to the growth chambers. A slice of fresh tissue was infiltrated, according to Jolivet et al. (1990), in a medium containing 100 mM mannitol, 10 mM HEPES, 10 mM MES (pH 6.6), and 0.2 mM CaCl2. Slice respiration was measured polarographically in the same medium, using a Clarke-type electrode (Rank Brothers Ltd., Cambridge, England). The alternative pathway was inhibited by adding 10 mM salicylhydroxamic acid (SHAM, resuspended in methoxy-ethanol) and the cytochrome pathway was inhibited by adding 1 mM potassium cyanide (KCN). Addition of SHAM and KCN together was used to record residual respiration (vres). Residual respiration was constant, around 6.1% of the total respiration, for both leaf and bulb tissues, regardless of the treatment or date of harvesting. In accordance with Bahr and Bonner (1973), it is postulated that the cytochrome pathway runs at a saturating rate under the experimental conditions (Vcyt=vcyt). Total respiratory rate (VT) was measured in the absence of inhibitors, whereas the activity of the cytochrome pathway (vcyt) was measured in the presence of SHAM as vcyt=VT+SHAM–vres, where VT+SHAM corresponds to the respiratory rate when SHAM was added first. The capacity of the alternative pathway (Valt) was measured in the presence of KCN as Valt=VT+KCN–vres., where VT+KCN represents the respiratory rate when KCN was added first. The activity of the alternative pathway (valt) was estimated as valt=VT–vcyt–vres. The engagement of the alternative pathway (ρ′) was determined as the ratio of valt to Valt. The participation of the alternative pathway (P) was determined as the ratio of valt to VT. Isolation of mitochondria Fresh bulbs (10 g) were cut in 100 ml of cold extraction medium containing 0.35 M mannitol, 30 mM MOPS buffer (pH 7.4), 7 mM cysteine, 2 mM EGTA, 2.5 mM MgCl2, 0.2% BSA (w/v), and 0.3% PVP (w/v). The bulbs were homogenized for 4× 1 s at full speed in a blender (Moulinex, Ecully, France). The homogenate was filtered through a 60 μm nylon net. The bulb fragments retained in the net were reblended in the mixer with 100 ml of the extraction medium. Homogenization and filtration were repeated as described above. The two successive filtrates were pooled and centrifuged according to Gerard and Dizengremel (1988). The mitochondrial pellet was resuspended in washing medium containing 0.35 mM mannitol, 10 mM MOPS buffer (pH 7.2), and 0.1% BSA w/v. Membrane integrity was measured spectrophotometrically at 550 nm, as the oxidation of reduced cytochrome c by washed intact versus burst mitochondria (Krippner et al., 1996). As mitochondrial isolation required much more material (10 g) than respiration on slices of tissue, it was only done for material harvested on day 16 for each treatment. Mitochondrial respiration Oxygen uptake was followed polarographically with a Clark-type electrode (Rank Brothers Ltd., Cambridge, England) on isolated mitochondria at 16 d. Respiratory studies were performed in a reaction medium containing 0.35 M mannitol, 5 mM MgCl2, 10 mm KC1, 0.1% BSA (w/v), and 10 mM phosphate buffer (pH 7.2). The oxidation of succinate (20 mM) was measured in the presence of 200 μM ATP at pH 7.2. The oxidation of malate (30 mM) was followed at pH 6.7 in MES buffer, and at pH 7.8 in TRIS-HCl buffer, as these two pHs were optimal for NAD-malic enzyme and malate dehydrogenase activity, respectively. The oxidation of malate at pH 7.8 was carried out in the presence of glutamate (2 mM) and NAD (400 μM) as a co-factor of malate dehydrogenase. The oxidation of NADH (1 mM) was followed at pH 7.2. Mitochondrial protein concentrations were determined according to Bradford (1976). KCN (1 mM) in aqueous solution and SHAM (700 μM) in methoxy-ethanol were used as inhibitors of the cytochrome and alternative pathways, respectively. VT, vcyt, valt, ρ′, and P were estimated in the same manner as for slice respiration measurements. Moreover, ADP/O ratio was estimated as the molar ratio of ADP added to the mitochondria to oxygen consumed following complete utilization of the ADP. This ratio represents the efficiency of oxidative phosphorylation. Respiratory control (RC) was determined as the ratio of state 3 (i.e. when ADP is in excess) to state 4 (i.e. when ADP is limiting) and represents the control exerted by ATP synthase on the electron transport chain. AOX immunoblot The amount of AOX was estimated by the Western blot method on the same isolated mitochondrial extracts used for respiration measurements. Mitochondrial proteins were extracted in 62.5 mM TRIS (pH 6.8) buffer containing 10% v/v glycerol, 2% v/v SDS, 0.005% v/v bromophenol blue, and 28 mM β-mercaptoethanol. The amount of total protein was fixed at 180 μg and separated by SDS-PAGE. Proteins were then transferred to a nitrocellulose membrane using 48 mM TRIS buffer containing 39 mM glycine, 0.04% SDS, and 20% v/v methanol. AOX monoclonal antibodies were used as primary antibodies and anti-mouse IgG fragments conjugated with peroxidase were used as secondary antibodies. The bands were revealed on the autoradiograms using SuperSignal Ultra Chemiluminescent Substrate. Densitometric analysis was performed to quantify the intensities of the bands corrected for the background using ImageJ software (NIH ImageJ, NIH, Bethesda, MD). Statistical analysis All variables were analysed by three-way ANOVA (Statistix 8.2, Analytical Software, Tallahassee, FL, USA) testing CO2 (Elevated versus Low), O3 (Elevated versus Low), and time (7–10 harvest dates depending on the variable) as fixed effects, except for mitochondrial respiration and AOX immunoblots for which several plants had to be pooled to get enough material for one measurement per treatment. In this case, no ANOVA could be performed and only technical repetitions were carried out. Each of the four treatments was assigned to one of the eight chambers, for a total of two repetitions per treatment (n=2 for most variables). A new series of plants were grown under the same conditions the following year. Plant growth variables were recorded each year to ensure the repeatability of growth conditions over the two years (n=4). Other variables were measured either in the first or the second year due to time constraints and the number of plants per chamber needed for each type of measurements. Significant effects were determined for P <0.05. A posteriori multiple comparisons tests were performed using Fisher's LSD. Pearson product–moment correlations (r) were also carried out on cumulative amount of C fixed per plant, Susy activity and bulb respiratory rate for each harvest date and each treatment. Results Net photosynthetic rate and leaf carboxylase activities Net photosynthetic rate increased quickly from leaf unfolding (5 d) to reach maximum rates at 9 d (Fig. 1), after which Pn decreased continuously until complete leaf senescence at day 28. When averaged over the whole life of the leaf, Pn under elevated CO2 was 59% higher than under control conditions, whereas Pn under elevated O3 was 29% lower, compared with the control (Table 1). Under elevated O3, elevated CO2 stimulated Pn by 25% until day 9, compared with the control. Subsequently, decreases in Pn under elevated CO2+O3 were faster than under ambient air. Thus, Pn values under elevated CO2+O3 no longer differed from those of the control after 15 d. Table 1. Results of three-way factorial ANOVA testing effects of elevated CO2 and elevated O3 treatments on E. americanum growth, gas exchanges, metabolites, enzyme activities and respiration over time CO2 O3 Time CO2×O3 CO2×time O3×time CO2×O3×time Leaf parameters     Net photosynthesis 121.19 *** 53.66 *** 45.85 *** 4.49 * 1.96 1.53 0.09     Rubisco activity 1.68 36.07 *** 11.54 *** 0.80 1.06 7.29 *** 1.05     PEPc activity 1.76 77.95 *** 26.75 *** 8.16 * 0.14 6.93 ** 1.11     Dry mass 0.03 0.09 20.50 *** 0.01 0.04 0.19 0.02     Sucrose 0.07 4.34 37.43 *** 0.35 0.16 * 2.24 0.41     Reducing sugars 0.12 0.40 0.93 0.25 0.13 0.30 0.50     Slice respiration 0.31 92.76 *** 426.16 *** 3.65 0.62 7.30 *** 0.77     Slice alternative respiration 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Slice alternative capacity 0.06 342.85 *** 451.84 *** 70.02 *** 5.40 ** 25.30 *** 5.87 **     Participation of the alternative pathway 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Engagement of the alternative pathway 2.41 30.77 *** – 0.43 – – – Bulb parameters     Dry mass 0.07 2.55 84.31 *** 0.22 0.12 0.29 0.33     Susy activity 38.43 *** 38.06 *** 84.97 *** 4.88 * 3.11 * 4.35 * 0.34     Invertase activity 0.15 4.29 375.35 *** 1.22 1.92 0.73 0.38     Starch 4.44 1.38 306.53 *** 0.00 0.62 1.02 0.21     Sucrose 2.19 0.17 165.57 *** 0.07 0.41 0.79 0.43     Reducing sugars 9.58 *** 6.30 * 112.01 *** 3.90 1.43 1.34 0.55     Slice respiration 359.49 *** 190.10 *** 843.79 *** 12.01 ** 23.56 *** 19.22 *** 19.30 ***     Slice alternative respiration 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Slice alternative capacity 905.09 *** 796.56 *** 522.52 *** 416.53 *** 78.97 *** 86.45 *** 51.32 ***     Participation of the alternative pathway 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Engagement of the alternative pathway 39.19 ** 57.42 ** – 8.24 – – – CO2 O3 Time CO2×O3 CO2×time O3×time CO2×O3×time Leaf parameters     Net photosynthesis 121.19 *** 53.66 *** 45.85 *** 4.49 * 1.96 1.53 0.09     Rubisco activity 1.68 36.07 *** 11.54 *** 0.80 1.06 7.29 *** 1.05     PEPc activity 1.76 77.95 *** 26.75 *** 8.16 * 0.14 6.93 ** 1.11     Dry mass 0.03 0.09 20.50 *** 0.01 0.04 0.19 0.02     Sucrose 0.07 4.34 37.43 *** 0.35 0.16 * 2.24 0.41     Reducing sugars 0.12 0.40 0.93 0.25 0.13 0.30 0.50     Slice respiration 0.31 92.76 *** 426.16 *** 3.65 0.62 7.30 *** 0.77     Slice alternative respiration 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Slice alternative capacity 0.06 342.85 *** 451.84 *** 70.02 *** 5.40 ** 25.30 *** 5.87 **     Participation of the alternative pathway 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Engagement of the alternative pathway 2.41 30.77 *** – 0.43 – – – Bulb parameters     Dry mass 0.07 2.55 84.31 *** 0.22 0.12 0.29 0.33     Susy activity 38.43 *** 38.06 *** 84.97 *** 4.88 * 3.11 * 4.35 * 0.34     Invertase activity 0.15 4.29 375.35 *** 1.22 1.92 0.73 0.38     Starch 4.44 1.38 306.53 *** 0.00 0.62 1.02 0.21     Sucrose 2.19 0.17 165.57 *** 0.07 0.41 0.79 0.43     Reducing sugars 9.58 *** 6.30 * 112.01 *** 3.90 1.43 1.34 0.55     Slice respiration 359.49 *** 190.10 *** 843.79 *** 12.01 ** 23.56 *** 19.22 *** 19.30 ***     Slice alternative respiration 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Slice alternative capacity 905.09 *** 796.56 *** 522.52 *** 416.53 *** 78.97 *** 86.45 *** 51.32 ***     Participation of the alternative pathway 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Engagement of the alternative pathway 39.19 ** 57.42 ** – 8.24 – – – F-values are presented along with statistical differences: * P <0.05, ** P <0.01, *** P <0.001. Open in new tab Table 1. Results of three-way factorial ANOVA testing effects of elevated CO2 and elevated O3 treatments on E. americanum growth, gas exchanges, metabolites, enzyme activities and respiration over time CO2 O3 Time CO2×O3 CO2×time O3×time CO2×O3×time Leaf parameters     Net photosynthesis 121.19 *** 53.66 *** 45.85 *** 4.49 * 1.96 1.53 0.09     Rubisco activity 1.68 36.07 *** 11.54 *** 0.80 1.06 7.29 *** 1.05     PEPc activity 1.76 77.95 *** 26.75 *** 8.16 * 0.14 6.93 ** 1.11     Dry mass 0.03 0.09 20.50 *** 0.01 0.04 0.19 0.02     Sucrose 0.07 4.34 37.43 *** 0.35 0.16 * 2.24 0.41     Reducing sugars 0.12 0.40 0.93 0.25 0.13 0.30 0.50     Slice respiration 0.31 92.76 *** 426.16 *** 3.65 0.62 7.30 *** 0.77     Slice alternative respiration 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Slice alternative capacity 0.06 342.85 *** 451.84 *** 70.02 *** 5.40 ** 25.30 *** 5.87 **     Participation of the alternative pathway 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Engagement of the alternative pathway 2.41 30.77 *** – 0.43 – – – Bulb parameters     Dry mass 0.07 2.55 84.31 *** 0.22 0.12 0.29 0.33     Susy activity 38.43 *** 38.06 *** 84.97 *** 4.88 * 3.11 * 4.35 * 0.34     Invertase activity 0.15 4.29 375.35 *** 1.22 1.92 0.73 0.38     Starch 4.44 1.38 306.53 *** 0.00 0.62 1.02 0.21     Sucrose 2.19 0.17 165.57 *** 0.07 0.41 0.79 0.43     Reducing sugars 9.58 *** 6.30 * 112.01 *** 3.90 1.43 1.34 0.55     Slice respiration 359.49 *** 190.10 *** 843.79 *** 12.01 ** 23.56 *** 19.22 *** 19.30 ***     Slice alternative respiration 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Slice alternative capacity 905.09 *** 796.56 *** 522.52 *** 416.53 *** 78.97 *** 86.45 *** 51.32 ***     Participation of the alternative pathway 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Engagement of the alternative pathway 39.19 ** 57.42 ** – 8.24 – – – CO2 O3 Time CO2×O3 CO2×time O3×time CO2×O3×time Leaf parameters     Net photosynthesis 121.19 *** 53.66 *** 45.85 *** 4.49 * 1.96 1.53 0.09     Rubisco activity 1.68 36.07 *** 11.54 *** 0.80 1.06 7.29 *** 1.05     PEPc activity 1.76 77.95 *** 26.75 *** 8.16 * 0.14 6.93 ** 1.11     Dry mass 0.03 0.09 20.50 *** 0.01 0.04 0.19 0.02     Sucrose 0.07 4.34 37.43 *** 0.35 0.16 * 2.24 0.41     Reducing sugars 0.12 0.40 0.93 0.25 0.13 0.30 0.50     Slice respiration 0.31 92.76 *** 426.16 *** 3.65 0.62 7.30 *** 0.77     Slice alternative respiration 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Slice alternative capacity 0.06 342.85 *** 451.84 *** 70.02 *** 5.40 ** 25.30 *** 5.87 **     Participation of the alternative pathway 3.87 152.43 *** 32.27 *** 13.47 ** 0.54 6.14 ** 0.10     Engagement of the alternative pathway 2.41 30.77 *** – 0.43 – – – Bulb parameters     Dry mass 0.07 2.55 84.31 *** 0.22 0.12 0.29 0.33     Susy activity 38.43 *** 38.06 *** 84.97 *** 4.88 * 3.11 * 4.35 * 0.34     Invertase activity 0.15 4.29 375.35 *** 1.22 1.92 0.73 0.38     Starch 4.44 1.38 306.53 *** 0.00 0.62 1.02 0.21     Sucrose 2.19 0.17 165.57 *** 0.07 0.41 0.79 0.43     Reducing sugars 9.58 *** 6.30 * 112.01 *** 3.90 1.43 1.34 0.55     Slice respiration 359.49 *** 190.10 *** 843.79 *** 12.01 ** 23.56 *** 19.22 *** 19.30 ***     Slice alternative respiration 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Slice alternative capacity 905.09 *** 796.56 *** 522.52 *** 416.53 *** 78.97 *** 86.45 *** 51.32 ***     Participation of the alternative pathway 305.05 *** 359.87 *** 24.44 *** 94.29 *** 15.70 *** 31.28 *** 20.11 ***     Engagement of the alternative pathway 39.19 ** 57.42 ** – 8.24 – – – F-values are presented along with statistical differences: * P <0.05, ** P <0.01, *** P <0.001. Open in new tab Fig. 1. Open in new tabDownload slide Time-courses of net photosynthetic rate (Pn) in leaves of E. americanum grown under control, elevated CO2, elevated O3, and elevated CO2+O3 during the epigeous growth period. The last data points (28 d) correspond to complete leaf senescence. The standard error, estimated from the MSE term in the ANOVA, is shown with the grand mean (N=2). The two vertical dotted lines indicate the end of leaf unfolding and the beginning of leaf senescence, respectively. Under ambient air, leaf rubisco activity increased rapidly until day 16 and then decreased before the first visual sign of leaf senescence, which occurs at day 21 (Fig. 2A). Leaf rubisco activity tended to increase more rapidly under elevated CO2 until day 12, but overall activity was not significantly different from that of the control (Table 1). O3 treatment decreased leaf Rubisco activity by 22% from day 12 to 20, regardless of CO2 concentration. However, leaf PEPc activity responded differently to O3 stimulation depending on CO2 concentration (Fig. 2B). Between days 12 and 20, O3-induced increases in PEPc activity were 56% and 38% under ambient and elevated CO2, respectively. Elevated CO2 concentrations alone did not affect leaf PEPc activity compared with the control. Fig. 2. Open in new tabDownload slide Time-courses of Rubisco carboxylase activity (A) and PEPc activity (B) in leaves of E. americanum grown under control, elevated CO2, elevated O3, and elevated CO2+O3 during the epigeous growth period The standard error, estimated from the MSE term in the ANOVA, is shown with the grand mean (N=2). The two vertical dotted lines indicate the end of leaf unfolding and the beginning of leaf senescence, respectively. Plant biomass Bulb biomass exhibited a large increase shortly after complete leaf unfolding (i.e. 5 d; Fig. 3). The maximum was reached at day 17, a few days before the initiation of leaf senescence (i.e. 21 d), after which time bulb biomass stopped increasing. No treatment significantly affected bulb growth kinetics or its final biomass, compared with the controls (Table 1). Bulb biomass represented 83±0.5% of total plant biomass at final harvest. Leaf and root biomass were fairly constant over time and similar among treatments. Fig. 3. Open in new tabDownload slide Time-courses of plant biomass of E. americanum grown under control (A), elevated CO2 (B), elevated O3 (C) and elevated CO2+O3 (D) during the epigeous growth period. The last data points (28 d) correspond to complete leaf senescence. Leaf, bulb, and roots are presented in black, white, and grey, respectively (N=4). The two vertical dotted lines indicate the end of leaf unfolding and the beginning of leaf senescence respectively. Bulb sucrose hydrolysis activities Invertase activity was high at the beginning of the season up to 8 d in all treatments, while Susy activity was minimum (Fig. 4). While invertase decreased progressively thereafter, Susy activity increased to maximum activity at 12 d. At 16 d, invertase activity reached minimum values around 20.1 nkat g−1 FW. Invertase activity was greater than Susy activity by 4-fold, suggesting that sucrose was mainly hydrolysed by invertase in the bulb of E. americanum (Fig. 4A). Elevated CO2 conditions increased Susy activity by 21% from day 8 to the initiation of senescence, compared with the control, whereas elevated O3 decreased Susy activity by 10% (Fig. 4B, Table 1). Susy activity was similar under elevated CO2+O3 and control conditions, and presented intermediate values between elevated CO2 and elevated O3 treatments. Elevated CO2 and elevated O3 did not affect invertase activity. Fig. 4. Open in new tabDownload slide Time-courses of cytoplasmic invertase activity (A) and sucrose synthase activity (B) in bulbs of E. americanum grown under control, elevated CO2, elevated O3, and elevated CO2+O3 during the epigeous growth period. The standard error, estimated from the MSE term in the ANOVA, is shown with the grand mean (N=2). The two vertical dotted lines indicate the end of leaf unfolding and the beginning of leaf senescence, respectively. Sugar accumulation Starch was absent from the leaf throughout the season in E. americanum. Sucrose concentrations in the leaf increased over time (Fig. 5A), whereas reducing sugar concentrations were constant, around 25 mg g−1 DW (Fig. 5B). Neither elevated CO2 nor elevated O3 treatments significantly affected leaf sucrose or reducing sugar concentrations (Table 1). Fig. 5. Open in new tabDownload slide Time-courses of sucrose (A, D), reducing sugar (B, E), and starch concentrations (C) in leaf (A, B), and bulb (C, D, E) of E. americanum grown under control, elevated CO2, elevated O3, and elevated CO2+O3 during the epigeous growth period. The standard error, estimated from the MSE term in the ANOVA, is shown with the grand mean (N=2). The two vertical dotted lines indicate the end of leaf unfolding and the beginning of leaf senescence, respectively. In the bulb, starch concentration exhibited a sigmoidal curve similar to that of the bulb mass, suggesting that the main constituent of bulb growth was starch accumulation. This result was confirmed by high final starch concentrations, which reached around 80% in the bulb, irrespective of the treatment (Fig. 5C). Starch concentration tended to increase more quickly after 4 d under elevated CO2 conditions than under the three other growth conditions (P=0.090). Neither elevated O3 nor elevated CO2+O3 conditions affected starch accumulation kinetics. After 16 d, starch concentrations were similar among treatments until leaf senescence. Sucrose exhibited a similar decrease in concentrations under all treatments (Fig. 5D). Similarly, reducing sugar concentrations decreased sharply after 4 d (Fig. 5E). However, elevated CO2 conditions maintained a 26% higher concentration from day 8 to complete leaf senescence, whereas elevated O3 conditions slightly reduced soluble sugar concentrations compared with the control. Leaf and bulb slice respiration In the leaf, total O2 consumption (VT) increased until 12–16 d and decreased abruptly several days before leaf senescence (Fig. 6A). Elevated CO2 curves were similar to the control curves (Table 1), whereas O3 treatment increased total leaf respiration by 18% between day 8 and day 16 (i.e. during the fastest growth phase), independent of CO2 concentration. During this period, leaf respiration inhibited by SHAM (valt) was 2-fold greater in O3-treated plants than in control plants, leading to an increased participation of the alternative pathway in total respiration (valt/VT), from 0.16 in the controls to 0.26 in both O3 treatments (Fig. 6B). Moreover, alternative pathway respiration tended to be higher at elevated CO2 concentrations than under control conditions, starting from day 8 (P=0.076). Elevated O3 increased the total capacity of the alternative pathway (Valt) by 59% compared with the controls (Figs 6C, 7A, C). Elevated CO2 did not affect the capacity of the alternative pathway (Fig. 7B). Engagement of the alternative pathway (valt/Valt) also increased under elevated O3 compared with the control. Fig. 6. Open in new tabDownload slide Time-courses of total respiratory rate (A, D), alternative pathway respiratory rate (B, E), and alternative pathway capacity (C, F) in leaf (A, B, C) and bulb (D, E, F) of E. americanum grown under control, elevated CO2, elevated O3, and elevated CO2+O3 during the epigeous growth period. Participation (B, E) and the engagement of the alternative pathway (C, F) are indicated on the graphs for day 16. The standard error, estimated from the MSE term in the ANOVA, is shown with the grand mean (N=2). The two vertical dotted lines indicate the end of leaf unfolding and the beginning of leaf senescence, respectively. Fig. 7. Open in new tabDownload slide Representative traces of respiratory activities of leaf (A, B, C, D) and bulb (E, F, G, H) slices from E. americanum grown under control (A, E), elevated CO2 (B, F), elevated O3 (C, G), and elevated CO2+O3 (D, H) during the epigeous growth period. The oxidation rates, expressed in nmol O2 min−1 mg−1 protein, are indicated on the different traces. In the bulb, total respiration increased during the first 12 d of growth, after which a maximum value was maintained up to the initiation of leaf senescence (Fig. 6D). Respiratory rates were 29% higher under elevated CO2 from day 12 onwards, whereas elevated O3 decreased respiration by 10%, compared with ambient air (Table 1). Elevated CO2+O3 conditions did not significantly affect respiratory rates. Bulb respiration inhibited by SHAM was up to 2.8 times higher under elevated CO2 conditions from day 12 onwards, compared with the control (Fig. 6E). This increase in alternative pathway respiration was responsible for 100% of the increase in total respiration under elevated CO2 conditions. Elevated O3 induced a 39% decrease in the activity of the alternative pathway, as averaged over day 12 to day 20, which was responsible for 81% of the decrease in respiratory rates under elevated O3. Thus, the participation of the bulb alternative pathway was increased from 0.18 in the control to 0.37 under elevated CO2 and decreased to 0.08 under elevated O3 at 16 d. Elevated CO2+O3 significantly affected neither alternative respiratory pathway rates nor alternative respiratory capacity in the bulb, compared with the control (Figs 6E, F, 7E, H). The alternative pathway capacity (Valt) in the bulb was 2.4 times higher under elevated CO2 compared with the control, whereas it was slightly lower than the control under elevated O3 (Figs 6F, 7F, G). Engagement of the alternative pathway increased from 0.84 in the control to 0.98 under elevated CO2 and was 2-fold lower under elevated O3, suggesting a full utilization of the AOX enzyme under elevated CO2. Engagement of the alternative pathway under elevated CO2+O3 conditions (0.80) was similar to that of the control. Bulb mitochondrial respiration Mitochondria were extracted from material harvested at 16 d, where the largest differences between treatments were observed for bulb slice respiration (Fig. 6). The integrity of washed mitochondrial membrane was high and constant among treatments (78–82%, data not shown). The respiratory rate of the bulb mitochondria was the highest in state 3 (i.e. when ADP is available in excess), with succinate as the initial substrate of the electron transport chain (i.e. 99 nmol O2 min−1 mg−1 protein; Fig. 8A), and lowest with NADH (i.e. 39 nmol O2 min−1 mg−1 protein; Fig. 8B). Malate oxidation was 37% higher when malate dehydrogenase activity was induced at pH 7.8 (81 nmol O2 min−1 mg−1 protein; Fig. 8C) than under malic enzyme activity at pH 6.7 (62 nmol O2 min−1 mg−1 protein; Fig. 8D). Succinate oxidation was partially inhibited by KCN (1 mM) and cyanide-insensitive respiration, which represented 18% of total respiration, was completely blocked by SHAM (700 μM). Similarly, cyanide-insensitive respiration represented 19% and 24% of the total respiration during malate oxidation at pH 7.8 and pH 6.7, respectively. These results suggest that electron flow was mediated by both the cytochrome and the alternative pathways for all three substrates. On the other hand, KCN almost completely blocked NADH oxidation by the electron transport chain. Fig. 8. Open in new tabDownload slide Representative traces of respiratory activities of washed mitochondria isolated from E. americanum bulbs grown under control (A, B, C, D), elevated CO2 (E, F), elevated O3 (G), and elevated CO2+O3 (H) during the epigeous growth period. Activities were recorded in presence of succinate (A), NADH (B) and malate (C–H) as substrates. Malate was oxidized in the presence of 2 mM glutamate and 400 μM NAD at pH 7.8 (C) and without glutamate at pH 6.7 (D–H). Respiratory activities with SHAM added first (E) and with KCN added first (F) were presented. Mitochondrial preparations used in each series of measures contained 200 μg of proteins. The oxidation rates, expressed in nmol O2 min−1 mg−1 protein, are indicated on the different traces. Malate oxidation at pH 6.7 exhibited treatment effects, on both respiratory rate and the alternative pathway, that were most similar to the treatment effects observed with slice respiration. In order to approximate the true physiological state and because of the central role played by malate in plant cell metabolism the behaviour of this substrate was therefore presented here to compare the four treatments. When SHAM was first added, it inhibited 15% of malate oxidation at pH 6.7 (i.e. mostly by malic enzyme), suggesting a rate (valt) of 9 nmol O2 min−1 mg−1 protein for the alternative pathway (Table 2). Elevated CO2 concentrations strongly stimulated malate oxidation from 68 nmol O2 to 91 nmol O2 min−1 mg−1 protein (VT). This stimulation was linked to a 4.2-fold higher activity of the cyanide-insensitive pathway (valt) and a 2.4-fold higher alternative pathway capacity (Valt). Elevated O3 affected malate oxidation at pH 6.7. The alternative pathway activity was slightly lower, whereas the alternative pathway capacity was reduced by 44% under elevated O3, compared with the controls. Elevated CO2+O3 conditions affected neither malate oxidation at pH 6.7 nor allocation of the mitochondrial electron flux to the alternative pathway or the cytochrome pathway. Moreover, as the activity of the cytochrome pathway was essentially constant under the different treatments (vcyt), modulation of malate oxidation at pH 6.7 can be entirely explained by changes in the activity of the alternative pathway. Thus, the engagement and participation of the alternative pathway was much higher under elevated CO2 compared with the control. Lower RC and a lower ADP/O ratio were recorded under elevated CO2 compared to the control, supporting the idea of increased participation of the alternative pathway. Indeed, the alternative pathway reduces the H+ gradient leading to reduced ATP synthesis. By contrast, RC increased under elevated O3. RC and the ADP/O ratio were also slightly higher under elevated CO2+O3 than under control conditions. Table 2. Respiratory activities and parameters of E. americanum bulb mitochondria isolated at 16 d from control and treated plants VT vcyt valt Valt ADP/O RC ρ′ P Control 68±6 58±4 9±3 16±6 2.34±0.3 1.48±0.4 0.57 0.13 Elevated CO2 91±4 53±1 38±4 39±3 1.98±0.2 1.07±0.2 0.97 0.42 Elevated O3 61±5 56±6 6±4 9±5 2.21±0.4 1.81±0.1 0.66 0.10 Elevated CO2+O3 69±11 59±8 11±6 16±7 2.67±0.3 1.69±0.2 0.69 0.16 VT vcyt valt Valt ADP/O RC ρ′ P Control 68±6 58±4 9±3 16±6 2.34±0.3 1.48±0.4 0.57 0.13 Elevated CO2 91±4 53±1 38±4 39±3 1.98±0.2 1.07±0.2 0.97 0.42 Elevated O3 61±5 56±6 6±4 9±5 2.21±0.4 1.81±0.1 0.66 0.10 Elevated CO2+O3 69±11 59±8 11±6 16±7 2.67±0.3 1.69±0.2 0.69 0.16 The substrate was malate (30 mM) at pH 6.7. Total respiratory rate (VT), the activity of the cytochrome pathway (vcyt), the activity of the alternative pathway (valt), the capacity of the alternative pathway (Valt), ADP/O ratio, the respiratory control (RC), the engagement of the alternative pathway (ρ′) and the participation of the alternative pathway (P) are presented (see Materials and methods for description of these variables). Values are the mean of three to four technical repetitions ±SE and expressed in nmol O2 min−1 mg−1 protein. Open in new tab Table 2. Respiratory activities and parameters of E. americanum bulb mitochondria isolated at 16 d from control and treated plants VT vcyt valt Valt ADP/O RC ρ′ P Control 68±6 58±4 9±3 16±6 2.34±0.3 1.48±0.4 0.57 0.13 Elevated CO2 91±4 53±1 38±4 39±3 1.98±0.2 1.07±0.2 0.97 0.42 Elevated O3 61±5 56±6 6±4 9±5 2.21±0.4 1.81±0.1 0.66 0.10 Elevated CO2+O3 69±11 59±8 11±6 16±7 2.67±0.3 1.69±0.2 0.69 0.16 VT vcyt valt Valt ADP/O RC ρ′ P Control 68±6 58±4 9±3 16±6 2.34±0.3 1.48±0.4 0.57 0.13 Elevated CO2 91±4 53±1 38±4 39±3 1.98±0.2 1.07±0.2 0.97 0.42 Elevated O3 61±5 56±6 6±4 9±5 2.21±0.4 1.81±0.1 0.66 0.10 Elevated CO2+O3 69±11 59±8 11±6 16±7 2.67±0.3 1.69±0.2 0.69 0.16 The substrate was malate (30 mM) at pH 6.7. Total respiratory rate (VT), the activity of the cytochrome pathway (vcyt), the activity of the alternative pathway (valt), the capacity of the alternative pathway (Valt), ADP/O ratio, the respiratory control (RC), the engagement of the alternative pathway (ρ′) and the participation of the alternative pathway (P) are presented (see Materials and methods for description of these variables). Values are the mean of three to four technical repetitions ±SE and expressed in nmol O2 min−1 mg−1 protein. Open in new tab Immunoblotted AOX protein Western blot analysis of mitochondria isolated from bulbs indicated that, at 16 d, the amount of AOX protein was 1.4 times higher under elevated CO2 (Fig. 9), and 3.4 times lower under elevated O3, compared with the control. Under elevated CO2+O3 conditions, AOX content was slightly lower than in the control plants. On the Western blots, AOX from E. americanum appeared as a monomeric reduced form with a molecular mass of 39 kDa. This apparent molecular mass of AOX coincided with the predicted molecular mass of the mature enzyme (39.331 kDa). Fig. 9. Open in new tabDownload slide A representative Western blot of AOX protein detection (above) and relative amount of AOX protein (below) in mitochondria isolated at 16 d from bulbs of E. americanum grown under control, elevated CO2, elevated O3, and elevated CO2+O3. Each value represents the mean of four technical repetitions ±SE. Discussion Modulation of the source strength Net photosynthetic rate of E. americanum was modulated by CO2 and O3 concentrations, as has been shown in numerous other species (Ceulemans and Mousseau, 1994; Ainsworth and Long, 2005; Vandermeiren et al., 2005). Pn was stimulated under elevated CO2 concentrations. It is well known that elevated CO2 conditions increase CO2 availability as a substrate for Rubisco, thereby positively affecting the carboxylation/oxygenation ratio. However, the enzyme itself appeared unaffected since the same rates were observed for in vivo Rubisco activity in elevated CO2 and in the control. By contrast, Rubisco activity was decreased under elevated O3. Such decreases have previously been observed and appear to be due to an alteration of Rubisco structure and expression (Pelloux et al., 2001). Elevated O3 stimulated leaf PEPc activity in E. americanum. This enzyme supplies the anaplerotic pathway with C skeletons (Dizengremel, 2001). Thus, stimulation of PEPc activity could compensate for C loss induced by Rubisco alteration under elevated O3. However, this compensation was partial since Pn were lower in O3-treated plants than in control plants. Thus, we succeeded in both stimulating and inhibiting source activity by increasing the CO2 and O3 concentrations, respectively. O3 stimulated leaf respiration in E. americanum. Eighty per cent of this increased respiratory rate under elevated O3 can be explained by the increased activity of the alternative pathway. Induction of AOX expression by O3 has previously been described and has been related to protective mechanisms avoiding ROS production in mitochondria (Tosti et al., 2006). Indeed, AOX protein prevents the over-reduction of the electron transport chain, especially the ubiquinone pool, thus alleviating ozone damage (Maxwell et al., 1999). The higher respiratory rate was partly responsible for the lower leaf Pn in O3-treated plants. On the other hand, elevated CO2 did not affect leaf respiration. Respiration is often inhibited under elevated CO2 concentrations, as demonstrated by Gonzalez-Meler et al. (1996). Moreover, these authors reported a direct inhibition of the cytochrome pathway by elevated CO2. This inhibition suggests that a portion of the electrons was transferred from the cytochrome pathway to the alternative pathway, which could explain the slightly higher activity of the alternative pathway observed in leaves of E. americanum under elevated CO2, but without any impact on total respiratory rates. Thus, leaf respiration seems to be modulated more by abiotic factors than by source or sink activity. Source–sink imbalance Neither starch nor soluble sugar accumulation in the leaf was observed over time under elevated CO2, nor a reduction in these constituents under elevated O3. Starch has often been described as a temporary storage sugar in the leaf which contributes to photosynthesis regulation (Goldschmidt and Huber, 1992). Twice-ambient CO2 often leads to an increase in leaf carbohydrates; for example, Long and Drake (1992) reported an increase of 52% for the soluble sugar content and 160% for starch content. Therefore, it can safely be assumed that more C was translocated to the bulb under elevated CO2, whereas the opposite occurred under elevated O3, compared with the controls, since the leaf did modulate its carbohydrate content in response to changes in Pn. Despite differences in the amount of C translocated to the bulb under the different growth conditions, biomass allocation patterns were similar among treatments. Biomass accumulation in the sink increases steadily during the epigeous growth phase, until it reached a maximum which precedes by a few days the first visual sign of leaf senescence. Numerous reports in the literature have been made regarding positive responses of plant growth to the elevated C supply under elevated CO2 conditions, including for perennial organs (Daymond et al., 1997; Donnelly et al., 2001). However, a lack of effect on plant growth is possible in sink-limited plants (Arp, 1991; Woodward, 2002). Thus, despite CO2 stimulation of the source activity, similar rates of biomass accumulation have been reported for onions from bulbing to bulb maturity (Daymond et al., 1997) and for potatoes before tuber initiation (Conn and Cochran, 2006). Thus, storage organ growth cannot increase in sink-limited conditions, whatever the strength of the C source. In these studies, the inability to use the surplus C in the sink organs leads to a down-regulation of Pn, which did not occur in E. americanum. Invertase was strongly activated at the beginning of bulb growth, whereas Susy became activated later in the season. Koch (2004) associated the activation of invertase with cell growth and division and Susy activation with cell differentiation and reserve accumulation. Similar sequential contributions of the different sucrose-cleaving enzymes, as a function of developmental stage, seem to take place in the bulb of E. americanum. During invertase activation (4–12 d), sink C-demand was high, as indicated by the kinetics of growth and starch accumulation in the bulb, which were maximum. This high demand could explain why neutral invertase activity was not affected by treatments. Sink limitation, i.e. growth and starch accumulation slow down, occurred later (from day 12 onward) when Susy activation took over. The rate of sucrose hydrolysis by Susy in the bulb was then stimulated by elevated CO2 concentrations and inhibited by O3 exposure. The activity of this enzyme was strongly correlated with the cumulative amount of CO2 fixed by the leaf (Fig. 10A). Among the main carbohydrates stored in the sink, only reducing sugar concentrations were modulated by CO2 and O3 concentrations, being higher under elevated CO2 and slightly lower under elevated O3. It therefore seems that modulation of Susy activity was sufficient to maintain a constant sucrose concentration in the bulb of E. americanum, despite the variable amount of carbohydrates being translocated to the bulb. Susy activity and expression are stimulated by sucrose availability, whereas invertase activity is regulated by the hexose pool (Geigenberger et al., 2004; Koch, 2004). Geigenberger et al. (2004), however, proposed a mechanism to drive carbon into starch synthesis and away from respiration in response to sucrose supply, based on redox-activation of ADP-glucose pyrophosphorylase by sucrose. In the present study, potential surpluses of sucrose under elevated CO2 did not affect starch accumulation or bulb growth of E. americanum. Thus, it appears that the higher amount of C fixed under elevated CO2 led to a higher flow of sucrose unloaded in the bulb, then to a higher concentration in glucose and fructose through a modulation of Susy. The opposite occurred when the amount of C fixed was lower, as was the case under elevated O3, thus avoiding in both cases, an accumulation of sucrose in the bulb. Our results suggest that modulation of the activity of Susy avoided AGPase activation and, instead, modulated the pool of glycolytic intermediates available for respiration. Yet it is also possible that AGPase was already maximal under the O3 treatment and that, therefore, the surplus of sucrose under all other growth conditions stimulated Susy activity and, sequentially, respiration. Fig. 10. Open in new tabDownload slide Relationship between the cumulative amount of carbon fixed by the leaves and sucrose synthase activity (A), respiratory rate of bulb slice (B), and the alternative respiratory rates of bulb slice (D); and between sucrose synthase activity and respiratory rate in bulb slices (C) of E. americanum. Control (black circles), elevated CO2 (black triangles), elevated O3 (white triangles), and elevated CO2+O3 plants (grey triangles). Pearson correlation coefficients and associated P-values are indicated. The role of the alternative respiratory pathway It seems that the bulb of E. americanum is unable to modulate its storage capacity in response to changes in the source activity, probably because growth in this species is sink-limited (Lapointe, 2001). Therefore, the different amounts of C translocated to the bulb must induce changes in C metabolism to counterbalance the constant growth rate of the perennial organ. O2 consumption by the bulb mitochondrial electron transport chain was stimulated under elevated CO2 and a reduction under elevated O3 conditions. Bulb respiratory rate is also strongly correlated with the cumulative amount of C fixed in the leaves (Fig. 10B) and with Susy activity (Fig. 10C). Shugaev and Sokolova (2001) reported a higher respiratory rate in non-differentiated stolons of potato where starch synthesis was low and glucose and fructose concentrations were high, compared with newly formed tubers. Storage organs seem to be able to use the respiratory process to remove the surplus of carbohydrates that cells cannot immediately store. Alternatively, reduced availability of sugars can induce a reduction in respiratory rates without affecting bulb growth rates, as seen under elevated O3. This result suggests that respiration was already stimulated under control conditions due to the incapacity of the sink to use all sugars for bulb growth. The use of inhibitors to estimate cytochrome and alternative pathway activities has been criticized (Millar et al., 1995), as it could underestimate the activity of the alternative pathway if the cytochromic pathway is not saturated before the inhibitors are added. Measurements of the capacity of the alternative pathway, on the other hand, are not subject to the same criticism. In the present study, the effects of the treatments on the alternative pathway activity were supported by similar effects on the alternative pathway capacity, in both bulb slices and mitochondrial extracts. Furthermore, the amount of AOX protein in the mitochondrial extracts varied in accordance with the activity and capacity of the alternative pathway. The development of a sink limitation during the season suggests a biological system almost saturated with carbohydrates. The difference between vcyt and Vcyt should therefore be minimum or nul. We are confident that, in the present study, the differences observed between treatments for the activity of the alternative pathway were not biased by the use of inhibitors. It appears that the increase in bulb respiration could be entirely explained by an increased activity of the alternative pathway in elevated CO2-grown plants. On the other hand, the decrease in respiration induced by the O3 treatment was partially due to a reduction in the activity of the alternative pathway (80%). This modulation of alternative pathway respiration was also strongly correlated with the cumulative amount of C fixed in the leaves (Fig. 10D). In isolated mitochondria, the activity of the alternative pathway also entirely explained the increase in the respiratory rate under elevated CO2 conditions in response to carbohydrate availability. Numerous studies have suggested that the alternative pathway could be a mechanism for energy overflow (Lambers, 1982), but few have tried to demonstrate it. The mitochondrial respiration data revealed a strong affinity of the alternative pathway for malate oxidation by NAD-malic enzyme in E. americanum bulbs. Moreover, this pathway increased under elevated CO2 conditions and decreased under elevated O3 conditions. NAD-malic enzyme may promote the reduction of the disulphide bond of the AOX protein, leading to the more active form of the enzyme (Vanlerberghe et al., 1995). Furthermore, pyruvate produced by NAD-malic enzyme is a well-known activator of the reduced AOX (Millar et al., 1993). Thus, NAD-malic enzyme activity and its subsequent organic acid production could play an important role in stimulating the consumption of excess carbohydrate by activation of the alternative respiratory pathway. In accordance with this hypothesis, both the capacity of the alternative pathway and the abundance of AOX were stimulated under elevated CO2, and repressed under elevated O3. Sieger et al. (2005) reported a modulation of AOX abundance and capacity in tobacco cells growing under limiting macronutrient conditions, and concluded that the alternative pathway was stimulated to correct the imbalance between carbohydrate supply and demand, thus controlling anabolism and growth. Stimulation of the alternative pathway would allow consumption of excess C, avoiding early senescence induced by a reduction in C sink demand. The capacity of the alternative pathway was strongly stimulated by CO2 and Western blot analysis also revealed a stimulation of AOX abundance, although of smaller amplitude. On the other hand, the capacity of the alternative pathway was slightly inhibited by ozone, whereas AOX abundance was strongly inhibited. Alternative oxidase exists as a covalent and a non-covalent dimer, the former being the less active form of the enzyme (Umbach and Siedow, 1993). The present results suggest that CO2 mainly promotes the proportion of the more active form (i.e. non-covalent dimer), leading to high capacity of the alternative pathway, whereas O3 mainly depletes the total amount of AOX protein. It has also been shown that only the more active form is susceptible to pyruvate activation (Umbach et al., 1994). This would agree with the strong stimulation of activity of the alternative pathway by elevated CO2 and of malate consumption by NAD-malic enzyme that was observed with the mitochondrial preparations. Both activation and expression of the AOX thus appear to be modulated as a function of C availability within the bulb. In this study, the response of sink C metabolism to the modulation of source activity was examined in E. americanum. Despite the increase in Pn under elevated CO2 and decrease under elevated O3, neither bulb growth nor starch accumulation was affected. Susy activity responds to the amount of sucrose translocated to the bulb. This modulation of the enzyme could prevent AGPase activation by sucrose content unless AGPase was already at a maximal rate. In addition, the bulb alternative pathway—activity, capacity, and AOX content—was modulated in response to the amount of C fixed. This modulation was stronger when malate was oxidized as the mitochondrial substrate by the malic enzyme. It is hypothesized that the production of pyruvate by malic enzyme modulates the activity of the AOX, allowing respiration to burn more C in excess conditions and less in more limited C conditions. The regulation of the glycolytic intermediate pool by Susy and of the alternative pathway by pyruvate production adjusts starch accumulation in rhythm with sink growth capacity. In this sink-limited species, sink C metabolism is modulated in response to changes in C availability, whereas source C metabolism mostly appeared to respond to growth conditions. It would be interesting to investigate the kinetics of starch synthesis and remobilization as a function of C availability and sink capacity to determine if other key enzymes are modulated by sink capacity. 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Rodríguez, Andrés Alberto; Maiale, Santiago Javier; Menéndez, Ana Bernardina; Ruiz, Oscar Adolfo

doi: 10.1093/jxb/erp256pmid: 19717530

Abstract The possible involvement of apoplastic reactive oxygen species produced by the oxidation of free polyamines in the leaf growth of salinized maize has been studied here. Salt treatment increased the apoplastic spermine and spermidine levels, mainly in the leaf blade elongation zone. The total activity of polyamine oxidase was up to 20-fold higher than that of the copper-containing amine oxidase. Measurements of H2O2, ·O2−, and HO· production in the presence or absence of the polyamine oxidase inhibitors 1,19-bis-(ethylamine)-5,10,15 triazanonadecane and 1,8-diamino-octane suggest that, in salinized plants, the oxidation of free apoplastic polyamines by polyamine oxidase by would be the main source of reactive oxygen species in the elongation zone of maize leaf blades. This effect is probably due to increased substrate availability. Incubation with 200 μM spermine doubled segment elongation, whereas the addition of 1,19-bis-(ethylamine)-5,10,15 triazanonadecane and 1,8-diamino-octane to 200 μM spermine attenuated and reversed the last effect, respectively. Similarly, the addition of MnCl2 (an ·O2− dismutating agent) or the HO· scavenger sodium benzoate along with spermine, annulled the elongating effect of the polyamine on the salinized segments. As a whole, the results obtained here demonstrated that, under salinity, polyamine oxidase activity provides a significant production of reactive oxygen species in the apoplast which contributes to 25–30% of the maize leaf blade elongation. Growth, maize, polyamine oxidase, polyamines, reactive oxygen species, salinity Introduction Reactive oxygen species (ROS), namely the superoxide radical (·O2−), the hydroxyl radical (HO·), and H2O2 are the major apoplastic ROS (aROS) in plants (Schopfer et al., 2001). aROS are necessary in many plant developmental processes (Foreman et al., 2003; Demidchik and Maathuis, 2007), particularly in the elongation zone (EZ) of maize leaves during leaf extension (Rodríguez et al., 2002). In these plants, the salt-induced decrease of aROS contributes to the reduction of leaf elongation (Rodríguez et al., 2004). On the other hand, the diminution of the aforementioned aROS has been attributed to the inhibitory effect of NaCl on the NADPH oxidase (NOX) complex (Rodríguez et al., 2007). It has been shown that non-enzymatic processes involving reactive oxygen species (ROS) cause wall polysaccharide scission in vitro (Miller, 1986; Fry, 1998; Schweikert et al., 2000; Fry et al., 2001) and in vivo (Schopfer, 2001). It has also been suggested that a delicate equilibrium between cleavage and cross-linking activities by ROS may take place in the apoplast (Cosgrove, 1999). Under optimal conditions, NOX is the main source of apoplastic ·O2− (Schopfer et al., 2001), which dismutes to H2O2 through superoxide dismutase (SOD) activity. Remarkably, despite the fact that NaCl inhibits NOX activity, plants continue producing aROS in low concentrations and growing at a reduced rate (Rodríguez et al., 2004). Up to now, the origin of those aROS is unknown, and whether such a low aROS amount may still contribute to plant growth under salt stress conditions is uncertain. Polyamines (PA) are small organic polycations, naturally found in eukaryotic and prokaryotic cells, which have been associated with cell growth and development (Bais et al., 1999; Steiner et al., 2007). In plant cells, the most abundant PA are putrescine (Put), spermidine (Spd), and spermine (Spm, Kaur-Sawhney et al., 2003). Although PA are detected in both symplastic and apoplastic compartments (Torrigiani et al., 1986; Pistocchi et al., 1988; Slocum, 1991; Tiburcio et al., 1997), their biosynthesis takes place only in symplastic subcellular localizations (Slocum, 1991; Borrell et al., 1995; Tiburcio et al., 1997). In turn, PA cross the plant cell membrane towards the apoplast via a still unknown mechanism (Cona et al., 2006a), where they are catabolized by amine oxidases (AOs), enzymes associated with apoplastic compartments (Federico and Angelini, 1991; Angelini et al., 1995; Tavladoraki et al., 1998; Cona et al., 2006a). The copper-containing amine oxidase (CuAO) catabolizes the oxidation of lower PA, such as Put and cadaverine, on primary amino groups, whereas plant polyamine oxidase (PAO) oxidizes higher PA, Spd, and Spm on their secondary amino groups (Federico and Angelini, 1991). PA oxidation produces hydrogen peroxide (H2O2) among other products (Lim et al., 2006). Interestingly, some reports have demonstrated that, unlike NOX, PAO activity is stimulated by NaCl in Brassica campestris (Das et al., 1995). Therefore, in the present work, the possibility is addressed that maize AOs maintain or even increase their activity under saline conditions, thus contributing to keep the basal ROS level needed to uphold leaf growth under saline stress. For this purpose, PA levels and AOs activities upon salinization were determined, as well as the effect of PA concentration on growth of the most actively elongating region of the salinized maize leaf. Materials and methods Plant material Maize seeds (Zea mays cv. Prozea 30, Produsem, Pergamino, Argentina) were sown on moist vermiculite contained in plastic net frames placed over 4.5 l black plastic trays with aerated water. Trays were kept at 25 °C under a light panel of fluorescent and incandescent light bulbs providing 95 μmol photons m−2 s−1 illumination, with a 16 h photoperiod. When the second leaf emerged, 6 d after sowing, the water was changed to half-strength Hoaglands solution (Hoagland and Arnon, 1950), which included 25 mM NaCl in the saline treatment. This solution was changed daily, increasing the NaCl concentration from 25 mM to 50 mM, 100 mM, and finally, 150 mM NaCl. Solutions were thereafter refreshed every 2 d. At harvest, 14 d after seeding, the region spanning 10–20 mm from the ligule was sectioned. This segment was used throughout the experiments. Segment elongation measurements The elongation of the leaf blade segments was measured as previously described by Rodríguez et al. (2004). Segments were gently vacuum infiltrated for 1.5 min and incubated for 7 h in different solutions. Digital images of segments were obtained before and after the incubation period, using a scanner (HP PSC 1510, Hewlett Packard Company, Palo Alto, CA). Segment length was measured with image processing software (Optimas 6.1, Optimas Corporation, Bothell, WA) and segment growth was expressed as a percentage of length increase, with respect to the control in that period. Extraction of free PA, 1,3-diaminopropane (Dap), and apoplastic Na+ To extract free PA from the cell extracts (cPA), 30 leaf blade segments were frozen in liquid N2 and homogenized. The homogenate (300 mg) was resuspended in 1 ml of PCA 5% (v/v), incubated in ice for 30 min and centrifuged at 15 000 g (15 min). The pellet was discarded and the supernatant was kept at –20 °C (solution A). Maize apoplastic fluid extraction was performed according to Rodríguez et al. (2002) with modifications. Segment pools were introduced within a net bag, which was, in turn, placed inside a plastic tube. To extract free PA and Dap from apoplastic extract (aPA and aDap, respectively), tubes were centrifuged for 10 min at 2000 g and the fraction collected was lyophilized, resuspended in 200 μl perchloric acid (PCA) 5% (v/v) (solution B) and used for free PA extraction according to Marina et al. (2008). For apoplastic Na+ extraction, plastic tubes were centrifuged for 1 min at 1000 g to discard the washing solution and centrifuged again for 10 min at 2000 g to collect the apoplastic fluid. All centrifugations were done at 4 °C and the apoplastic fluids obtained were kept at –20 °C. Thirty segments from 30 leaves were used for free aPA and aDap extractions, and 120 segments from 24 leaves for Na+ extraction. In addition, 30 leaf blade segments from 10 unsalinized plants were pooled, washed, transferred to net bags, and gently vacuum infiltrated for 2 min with water or 100 mM NaCl. The resulting apoplastic fluid was used to determine the apoplastic peroxidase (POX) levels and to check for variations in free aPA contents derived from the presence of salt in the apoplastic environment. Glucose 6-phosphate dehydrogenase activity, a marker of cytosolic contamination, was determined in every apoplastic fluid fraction according to Rodríguez et al. (2002). Determination of free PA and Dap Maize free PA were determined according to Jiménez-Bremont et al. (2007). For dansylation, 200 μl of solution A or B (see above) were added to 10 μl of 0.1 mM heptanodiamine (internal standard, ICN) plus 200 μl saturated Na2CO3 and 400 μl dansyl chloride-acetone 1% (w/v). After 16 h at 25 °C in the dark, 100 μl of proline 100% (w/v) was added to stop the reaction. Dansyl-derived PA were extracted with 500 μl toluene. The organic phase (400 μl) was evaporated under vacuum and resuspended in 400 (cPA) or 200 (aPA and aDap) μl acetonitrile. Dansyl-derived PA were separated by HPLC (ISCO 2350, ISCO Inc, Lincoln, NE) with a reverse phase column Sephasil C18 (Amersham Pharmacia) and detected with a spectrofluorometer (Variant Fluorichrom). The solvent mix was obtained with a gradient programmer ISCO 2360, with a flow of 1.5 ml min−1 as follows: 0–4.5 min, acetonitrile:H2O 70:30 v/v; 4.5–9 min, acetonitrile 100; 9–15 min, acetonitrile:H2O 70:30 v/v). Peak areas were integrated, normalized to heptanodiamine and interpolated into a PA standards calibration curve. POX enzyme level The reaction mixture (1 ml) contained 15 μl of apoplastic fluid, 20 μl 0.02 M guaiacol, and 0.1 M potassium phosphate pH 6.4. The reaction was started by adding 35 μl 88 mM H2O2 and activity was measured as an increase in A560 after 30 s with a spectrophotometer (Beckman DU Series 600, Beckman Instruments, Fullerton, CA). The specific activity calculation was based on the protein content of each sample, determined according to Bradford (1976). Determination of the apoplastic Na+ content Apoplastic Na+ concentration was determined by atomic emission spectrophotometry analysis of the apoplastic fluid fraction, using a Perkin-Elmer AA 100 spectrophotometer in emission mode. H2O2 production by amine oxidase activity The AO activity level was determined according to Cona et al. (2006b) with some modifications. Segments were washed in water (control) or 100 mM NaCl (salinized) for 6 min in order to remove symplastic contamination. For in vivo measurements, pools of five segments were introduced in 1 ml solutions containing 100 μM 4-aminoantipyrine (4-AAP), 1 mM 3,5-dichloro-2-hydroxybenzenesulphonic acid (DCHBS), 20 mM potassium phosphate pH 6.5, plus or minus 0.5 mM Spm or Put, and 100 mM NaCl for the saline treatment. Segments were subsequently infiltrated for 2 min and further incubated for 5 h at room temperature. Then 1 ml of the incubation medium was collected and the resultant pink adduct was measured at A515 with a spectrophotometer (HITACHI U-2000, Hitachi, Tokyo, Japan) and transformed into an H2O2 molar concentration with a molar extinction coefficient at 515 nm (2.6×104 M−1 cm−1). PAO and CuAO activities were calculated as the difference in H2O2 produced between treatments containing and lacking substrate. Extraction of apoplastic PAO Extraction of apoplastic proteins was performed as described by Li (1993) with slight modifications (Maiale et al., 2008). Plant material (40 g) was cut in 2 mm pieces, washed in distilled water and vacuum-infiltrated with 100 ml 5 mM potassium phosphate pH 6.5 added with 200 mM NaCl. The vacuum was broken and re-established every 5 min, for three successive times. The apoplastic fluid was collected, cooled at 4 °C and added with 1 vol. of pre-cooled (–20 °C) Me2CO. The resulting solution was incubated at 4 °C, for 30 min and centrifuged at 15 000 g for 15 min. The supernatant was discarded and the pellet resuspended in 20 mM bis-tris-propane buffer pH 6.5 and applied to a DEAE-Sephacell column (1× 2 cm) equilibrated with the same solution. The eluted solution (Solution D) was kept at 4 °C until used. In vitro PAO activity assay For in vitro PAO activity measurement, pools of 20 segments were frozen with liquid N2, homogenized in 1 ml of 0.1 M potassium phosphate pH 6.5 at 4 °C, and centrifuged at 15 000 g for 15 min. The pellet was discarded and the homogenate was kept at 4 °C (sSolution C). PAO activity was determined according to Cona et al. (2006b). Previous tests determined that the optimal pH for PAO activities was 6.5. The reaction mixture contained 1 ml, 50 μl solution C or 80 μl D (see above) plus 100 μM 4-AAP, 1 mM DCHBS, 0.06 mg ml−1 horseradish POX, and 100 mM potassium phosphate pH 6.5. The mixture was incubated at 30 °C for 2 min. The reaction was started by adding 5 μl of 10 mM Spm and the activity was measured for 1 min at A515 with a spectrophotometer and transformed into an H2O2 molar concentration. PAO Ki for SL-11061 was calculated (Lineweaver and Burk, 1934). The Km value obtained for this preparation was Km=17.7 μM. Extraction and purification of plasma membrane for NOX activity determination Plasma membrane was prepared according to Larsson (1985) with some variations. Leaves (70 g) from 7-d-old plants were homogenized with an omnimixer by giving three 20 s pulses at full speed. The extraction solution (200 ml) contained 50 mM TRIS-HCl pH 7.5, 0.33 M sucrose, 1 mM EDTA, 0.1 mM MgCl2, 1 mM ascorbate, 1 mM DTT, 1 mM phenylmethylsulphonyl fluoride, and 0.6% (w/v) polyvinylpoly pyrrolidone. The homogenate was filtered through four layers of cheesecloth, and the filtrate centrifuged at 10 000 g for 10 min. Microsomes were pelleted from the supernatant by centrifugation at 140 000 g for 45 min and resuspended in 10 ml 5 mM potassium phosphate pH 7.8 containing 0.33 M sucrose and 3 mM KCl. The suspension was fractionated by the aqueous two-phase partitioning method (Larsson, 1985). Phase separations were carried out in a series of 10 g phase systems with a final composition of 6.2% (w/w) dextran T500 (Sigma), 6.2% (w/w) polyethylene glycol 3350 (Sigma), 0.33 M sucrose, 5 mM potassium phosphate pH 7.8, and 3 mM KCl. Three successive partitioning rounds yielded an upper phase (U3) and a lower phase (L3). U3 was 3-fold diluted in 10 mM TRIS-HCl buffer (pH 7.4) containing 0.33 M sucrose. The solution was centrifuged at 140 000 g for 60 min and the resulting pellet resuspended in 2 ml 10 mM TRIS-HCl buffer pH 7.4 and 0.33 M sucrose. All procedures were carried out at 4 °C. The enrichment in plasma membranes of the upper phase was monitored by the percentage of V-ATPase inhibition (Serrano, 1978). U3 was enriched in plasma membrane up to 90%. U3 was kept at 4 °C and used for enzyme activity immediately. NADPH oxidase activity NADPH oxidase activity was assayed spectrophotometrically according to Sagi and Fluhr (2001). The reaction medium contained 50 μl U3, 0.3 mM Na,3′-[1-[(phenylamino)-carbonyl]-3,4-tetrazolium](4-methoxy-6-nitro) benzenesulphonic acid hydrate (XTT) and 0.2 mM NADPH in 50 mM TRIS-HCl buffer pH 7.4. The reaction was initiated by adding NADPH. Data were transformed into an ·O2− molar extinction coefficient at 470 nm (2.16×104 M−1 cm−1). H2O2 production by NADH-dependent POX Reactions were carried out in 0.1 M potassium phosphate pH 4.5 containing 3 μg ml−1 horseradish peroxidase and 0.2 mM NADH (Frahry and Schopfer, 1998) with some modifications. Reactions were initiated by adding NADH and, 5 min later, aliquots of 500 μl were removed from the reaction mixture. NADH was eliminated with 0.1 M HCl followed by 0.1 M NaOH. H2O2 was measured by fluorescence of 55 μM homovanillic acid at 407 nm (EM) and 305 nm (EX), in the presence of 12 μg ml−1 horseradish peroxidase and 0.2 M potassium phosphate pH 4.5 in a final 1 ml volume. The calibration curve was linear in the range of 0.5–3 μM H2O2. The calibration curve was not affected by 100 μM or 200 μM SL-11061, 50 μM or 200 μM DPI, and 100 μM or 200 mM NaCl. Detection of ·O2− accumulation in the whole leaf ·O2− accumulation was detected by blue formazan precipitation (Hernández et al., 2001). For this purpose, 0.01% (w/v) nitro-blue tetrazolium (NBT) was added with nutrient solution to control and salinized 13-d-old plants. One day later, plants were harvested and the third leaf was boiled in 80% (v/v) ethanol for 10 min. Leaves were mounted on a glass slide and scanned. In vivo ·O2− production In blade segments, release of ·O2− to the medium was determined through spectrophotometry, using XTT (Frahry and Schopfer, 2001). Pools of eight segments were gently vacuum infiltrated and incubated 7 h in 1 ml of aqueous solutions containing 0.5 mM XTT, 100 mM NaCl and the following potential PAO activity modulators: 100–800 μM Spm, 200 μM SL-11061: 200 μM 1,8-diamino-octane (1,8-DO): 200 mM Dap, 1 mM ferrozine (FZ), and 1 mM neocuproine (NC). Segments were removed and the medium centrifuged at 10 000 g for 10 min. 1 ml of the incubation medium was collected, measured with a spectrophotometer at A470 and data transformed into an· ·O2−molar extinction coefficient at 470 nm (2.16×104 M−1 cm−1). In vivo HO· production HO· release to the medium was determined by the hydroxylation of sodium benzoate (BZ) by HO·. Hydroxyl BZ was detected by spectrofluorometry according to Schopfer et al. (2001) with modifications. Pools of six salinized segments were gently infiltrated and incubated for 7 h in 1 ml of aqueous solutions containing 2.5 mM BZ and 100 mM NaCl in the presence or absence of 100 μM SL-11061 at 30 °C in the dark. Fluorescence was determined at 407 nm emission after excitation at 305 nm in a spectrofluorometer (Bio-Tek Kontron SFM 25, Kontron Instruments, Zürich, Switzerland). Statistical analysis Data were analysed by one-way or two-way ANOVA and Tukey or DGC tests (Di Rienzo et al., 2002), using InfoStat (InfoStat 2007. InfoStat Group. Facultad de Ciencias Agropecuarias. Universidad Nacional de Córdoba. Version 1.1. Córdoba, Argentina). Results Effect of NaCl on elongation and apoplastic Na+ concentration and free PA levels in leaf segments Previous results showed that elongation in unsalinized and salt-treated leaves is maximal at the second blade EZ segment, that is, the region spanning 10–20 mm from the ligule (Fig. 1A). Elongation of excised second blade segments from salt-treated plants incubated in 100 mM NaCl was 50% compared with unsalinized segments (Fig. 1B), confirming the previous results by Rodríguez et al. (2004). Atomic emission spectrophotometry (AES) analysis of the segment apoplastic fluids revealed a 76.4±2.4 and 1.4±0.5 mM Na+ content in salinized and unsalinized leaf blades, respectively (no cytosolic contamination was detected in the apoplastic fluid; see Supplementary Table S1 at JXB online). Therefore, as the saline content of the incubation solution was comparable with that existing in the apoplast of salinized leaves, it was decided to add 100 mM NaCl to the incubation mixture in the next in vivo experiments, in the case of salinized plants, as a means to avoid changes in the osmotic potential of the apoplastic environment. Free PA levels were measured by HPLC. Salinity lowered Put and increased Spd and Spm level of the cell extracts (Fig. 2A–C), whereas it greatly increased Spm and Spd and slightly increased Put in the apoplast (Fig. 2D–F), suggesting a role for PA accumulation in the elongation zone of the maize leaf blade under saline stress. Fig. 1. Open in new tabDownload slide Schematic representation of expanding and expanded regions in unsalinized and salt-treated maize leaf blades and the effect of NaCl on the elongation of second blade EZ segments. (A) Distribution of EZ and maturation zone (MZ) in unsalinized and salt-treated maize leaf blades, adapted from Rodríguez et al. (2004). (B) Effect of NaCl on the elongation of second blade EZ segments. Segments were incubated for 7 h in water (unsalinized) or in solutions containing 100 mM NaCl (salt-treated). Segments were scanned before and after the incubation period and their length measured with an image processing software. Results are the percentage of length increase during a 7 h incubation period with respect to the control unsalinized segment. Absolute growth rate for unsalinized second blade segments was 0.402±0.011 mm h−1. The experiment was conducted twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=20). Asterisks indicate a difference from the control (P <0.05). Fig. 1. Open in new tabDownload slide Schematic representation of expanding and expanded regions in unsalinized and salt-treated maize leaf blades and the effect of NaCl on the elongation of second blade EZ segments. (A) Distribution of EZ and maturation zone (MZ) in unsalinized and salt-treated maize leaf blades, adapted from Rodríguez et al. (2004). (B) Effect of NaCl on the elongation of second blade EZ segments. Segments were incubated for 7 h in water (unsalinized) or in solutions containing 100 mM NaCl (salt-treated). Segments were scanned before and after the incubation period and their length measured with an image processing software. Results are the percentage of length increase during a 7 h incubation period with respect to the control unsalinized segment. Absolute growth rate for unsalinized second blade segments was 0.402±0.011 mm h−1. The experiment was conducted twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=20). Asterisks indicate a difference from the control (P <0.05). Fig. 2. Open in new tabDownload slide Free PA levels in segments of unsalinized and salt-treated plants. (A–C) Pools of second blade segments were homogenized and cell extracts used for cPA measurements. (D–F) aPA were extracted from second blade segment pools by centrifugation and the collection of apoplastic fluid. PA were dansyl-derived according to Jiménez-Bremont et al. (2007) and determined by HPLC. The experiment was performed twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=6). Asterisks indicate a difference from the control (P <0.05). Fig. 2. Open in new tabDownload slide Free PA levels in segments of unsalinized and salt-treated plants. (A–C) Pools of second blade segments were homogenized and cell extracts used for cPA measurements. (D–F) aPA were extracted from second blade segment pools by centrifugation and the collection of apoplastic fluid. PA were dansyl-derived according to Jiménez-Bremont et al. (2007) and determined by HPLC. The experiment was performed twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=6). Asterisks indicate a difference from the control (P <0.05). Evaluation of the mechanisms involved in aPA increment in salinized leaves It has been hypothesized that after being synthesized in the cytoplasm, PA cross the plasma membrane towards the apoplast, where they are catabolized by AOs. Thus, the observed rise in free aPA in salinized plants is probably the result of: (i) promotion of PA passage towards the apoplast or (ii) a decrease in the amount of AOs enzymes (or AOs activities) leading to free aPA accumulation. Given that the mechanisms of PA passage to the apoplast are unknown, it was decided to assess the last possibility. For this, the effect of salt addition on maximal in vivo and in vitro AO activity was determined. In this approach, H2O2 production by PA oxidation was estimated through an oxidative POX-dependent reaction that produces a pink adduct measurable by spectrophotometry (Cona et al., 2006b). Blade segments from salinized and unsalinized plants were infiltrated and incubated in the reaction mix with the addition of 0.5 mM of exogenous substrates Put and Spm, for CuAO and PAO determination, respectively (POX addition to the reaction mixture was not necessary since no variation was observed in the apoplastic oxidative POX activity between treatments, see Supplementary Fig. S1 at JXB online). As result, it was observed that (i) there were no differences in the maximal CuAO and PAO activities (achieved under saturating substrate conditions) due to salt treatment (Fig. 3A), (ii) PAO activity levels were up to 20-fold higher than those of CuAO (therefore further studies will be performed only on PAO activity). A second in vitro analysis using cell-free extracts from segment homogenates (and saturating substrate conditions) confirmed former results on PAO activity levels (Fig. 3B). Taken together, these results led us to: (i) reject the possibility of a negative saline effect on the total activities of AOs enzymes and (ii) to assume that salinity somehow promoted PA passage from the symplastic compartment towards the apoplast. Having in mind that aPA may interact with cell wall components, a third possibility is that the presence of NaCl in the apoplast causes the dissociation of pre-existent aPA from the cell wall. Blade segments from unsalinized plants were then infiltrated either with water or with 100 mM NaCl, resulting in the absence of any effect of this salt on free aPA contents (see Supplementary Table S2 at JXB online). Thus, any dissociating action of NaCl on aPA putatively associated to cell wall components of the apoplast was ruled out. Fig. 3. Open in new tabDownload slide PAO and CuAO activities under substrate saturating conditions. (A) In vivo AOs activities were determined according to Cona et al. (2006b). Pools of five segments were introduced in 1 ml of solution containing 100 μM 4-AAP, 1 mM DCHBS, 20 mM potassium phosphate pH 6.5 with and without 0.5 mM Spm or Put, and 100 mM NaCl for salinized segments. Segments were subsequently infiltrated for 5 min, incubated for 5 h and AOs activities determined by pink adduct production at A515. Data were transformed into H2O2 molar concentrations with a molar extinction coefficient at 515 nm (2.6×104 M−1 cm−1). PAO and CuAO levels were calculated as the difference in H2O2 amounts between treatments with and without substrate. (B) In vitro PAO measurement. Pools of 20 blade segments were homogenized in 1 ml of 0.1 mM potassium phosphate pH 6.5 at 4 °C, and centrifuged at 15 000 g for 15 min. PAO activity was determined according to Cona et al. (2006b). The experiment was conducted twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=6). Fig. 3. Open in new tabDownload slide PAO and CuAO activities under substrate saturating conditions. (A) In vivo AOs activities were determined according to Cona et al. (2006b). Pools of five segments were introduced in 1 ml of solution containing 100 μM 4-AAP, 1 mM DCHBS, 20 mM potassium phosphate pH 6.5 with and without 0.5 mM Spm or Put, and 100 mM NaCl for salinized segments. Segments were subsequently infiltrated for 5 min, incubated for 5 h and AOs activities determined by pink adduct production at A515. Data were transformed into H2O2 molar concentrations with a molar extinction coefficient at 515 nm (2.6×104 M−1 cm−1). PAO and CuAO levels were calculated as the difference in H2O2 amounts between treatments with and without substrate. (B) In vitro PAO measurement. Pools of 20 blade segments were homogenized in 1 ml of 0.1 mM potassium phosphate pH 6.5 at 4 °C, and centrifuged at 15 000 g for 15 min. PAO activity was determined according to Cona et al. (2006b). The experiment was conducted twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=6). NaCl increases inherent PAO activity So far, it has been demonstrated that aPA levels increased as a result of plant salinization (Fig. 2). This result encouraged us to examine whether the inherent PAO activity, which depends on the concentration of its endogenous substrate (aPA), correlates with that result. Therefore, PAO activity was evaluated by measuring the Dap content using HPLC. Dap is a product of the Spd and Spm oxidation, formed in the same molar quantities as H2O2 (Cona et al., 2006a). The measurement of aDap levels indicated that salinity led to increased PAO activity in the leaf blade region under study (Fig. 4A). Alternatively, PAO activity in vivo was determined as previously described (Fig. 3A), without the exogenous Spm supplement. For this purpose, H2O2 levels were measured in the presence or absence of 1,19-bis-(ethylamine)-5,10,15 triazanonadecane (SL-11061), a tobacco PAO inhibitor (Marina et al., 2008), which has also been found to inhibit oat PAO in vivo (Maiale et al., 2008). The results showed that SL-11061 had no effect on apoplastic H2O2 content in the absence of NaCl (Fig. 4B), showing that the contribution of PAO activity to the total apoplastic H2O2 level was negligible under control conditions. Conversely, a 50% lowered H2O2 content was found in SL-11061-treated segments under saline conditions. As a whole, these results showed that, under salt stress conditions, the contribution of PAO to the observed apoplastic H2O2 pool in the elongation zone of the maize leaf blade is relevant, whereas in the absence of salt treatment, the formation of the main apoplastic H2O2 would rely on mechanisms different from aPA oxidation. Fig. 4. Open in new tabDownload slide Effect of salinity on PAO activity without exogenous substrate. (A) aDap was determined in apoplastic fluids obtained from pools of 30 second segments. Dap dansylation was performed according to Jiménez-Bremont et al. (2007) and dansyl-derived Dap was determined through HPLC. (B) H2O2 production in segments from unsalinized and salt-treated plants was measured as in Fig. 3A, although without exogenous Spm, after 7 h incubation in presence or absence of 100 μM SL-11061. The experiment was performed twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=4). Bars sharing the same letter are not significantly different (P <0.05). Fig. 4. Open in new tabDownload slide Effect of salinity on PAO activity without exogenous substrate. (A) aDap was determined in apoplastic fluids obtained from pools of 30 second segments. Dap dansylation was performed according to Jiménez-Bremont et al. (2007) and dansyl-derived Dap was determined through HPLC. (B) H2O2 production in segments from unsalinized and salt-treated plants was measured as in Fig. 3A, although without exogenous Spm, after 7 h incubation in presence or absence of 100 μM SL-11061. The experiment was performed twice, yielding similar results. Abbreviations: c, unsalinized; s, salt-treated. Results are means ±SE (n=4). Bars sharing the same letter are not significantly different (P <0.05). On the other hand, under in vitro conditions NaCl did not affect PAO activity at all (Table 1), reinforcing that this enzyme keeps its activity in salinity. In addition, a low Ki value (8.7×10−7 M) was found for the polyamine analogue SL-11061, indicating its high efficiency as a maize PAO inhibitor. This Ki is comparable to that found for oat PAO (Ki=1.5×10−9 M; Maiale et al., 2008). Table 1. Effects of inhibitors on apoplastic ROS-producing enzymes Inhibitor treatment PAO NOX NADH-dependent POX (% of control) (% of control) Control 100.0±0.3 100.0±6.7 100.0±3.1 SL-11061 (μM)     5 31.2±0.8* – –     20 9.9±0.3* – –     100 0.0±0.0* 105.5±1.0 106.3±11.0     200 0.0±0.0* 104.8±0.1 102.5±6.2 DPI (μM)     20 99.7±2.7 23.3±0.5*a 6.0±6.0*b     200 95.8±0.1* 8.0±0.1* – NaCl (mM)     100 101.1±0.2 37.8±1.6*a 41.5±4.6*     200 102.3±0.1 10.7±0.6* 28.9±0.6* Inhibitor treatment PAO NOX NADH-dependent POX (% of control) (% of control) Control 100.0±0.3 100.0±6.7 100.0±3.1 SL-11061 (μM)     5 31.2±0.8* – –     20 9.9±0.3* – –     100 0.0±0.0* 105.5±1.0 106.3±11.0     200 0.0±0.0* 104.8±0.1 102.5±6.2 DPI (μM)     20 99.7±2.7 23.3±0.5*a 6.0±6.0*b     200 95.8±0.1* 8.0±0.1* – NaCl (mM)     100 101.1±0.2 37.8±1.6*a 41.5±4.6*     200 102.3±0.1 10.7±0.6* 28.9±0.6* PAO was extracted and purified from apoplastic fluids of leaf blade segments according to Li (1993) with some modifications. PAO activity was determined according to Cona et al. (2006b). NOX was extracted and purified according to Larsson (1985) with some variations. NOX activity was determined according to Sagi and Fluhr (2001). H2O2 production by NADH-dependent POX was determined according Frahry and Schopfer (1998) with modifications. Data are means ±SE (n=3). Asterisks indicate significantly different from control (P <0.05). 100% of control for PAO, NADH-dependent POX and NOX represent: 6.15±0.02 μmol H2O2 min−1, 389±9 μmol H2O2 min−1 g−1, and 20.05±0.62 pmol ·O2− min−1, respectively. PAO, NOX, and NADH-dependent POX substrate concentrations (Spm, NADH, and NADPH, respectively) were 200 μM in all cases. a From Rodríguez et al. (2007). b From Frahry and Schopfer et al. (2007). Open in new tab Table 1. Effects of inhibitors on apoplastic ROS-producing enzymes Inhibitor treatment PAO NOX NADH-dependent POX (% of control) (% of control) Control 100.0±0.3 100.0±6.7 100.0±3.1 SL-11061 (μM)     5 31.2±0.8* – –     20 9.9±0.3* – –     100 0.0±0.0* 105.5±1.0 106.3±11.0     200 0.0±0.0* 104.8±0.1 102.5±6.2 DPI (μM)     20 99.7±2.7 23.3±0.5*a 6.0±6.0*b     200 95.8±0.1* 8.0±0.1* – NaCl (mM)     100 101.1±0.2 37.8±1.6*a 41.5±4.6*     200 102.3±0.1 10.7±0.6* 28.9±0.6* Inhibitor treatment PAO NOX NADH-dependent POX (% of control) (% of control) Control 100.0±0.3 100.0±6.7 100.0±3.1 SL-11061 (μM)     5 31.2±0.8* – –     20 9.9±0.3* – –     100 0.0±0.0* 105.5±1.0 106.3±11.0     200 0.0±0.0* 104.8±0.1 102.5±6.2 DPI (μM)     20 99.7±2.7 23.3±0.5*a 6.0±6.0*b     200 95.8±0.1* 8.0±0.1* – NaCl (mM)     100 101.1±0.2 37.8±1.6*a 41.5±4.6*     200 102.3±0.1 10.7±0.6* 28.9±0.6* PAO was extracted and purified from apoplastic fluids of leaf blade segments according to Li (1993) with some modifications. PAO activity was determined according to Cona et al. (2006b). NOX was extracted and purified according to Larsson (1985) with some variations. NOX activity was determined according to Sagi and Fluhr (2001). H2O2 production by NADH-dependent POX was determined according Frahry and Schopfer (1998) with modifications. Data are means ±SE (n=3). Asterisks indicate significantly different from control (P <0.05). 100% of control for PAO, NADH-dependent POX and NOX represent: 6.15±0.02 μmol H2O2 min−1, 389±9 μmol H2O2 min−1 g−1, and 20.05±0.62 pmol ·O2− min−1, respectively. PAO, NOX, and NADH-dependent POX substrate concentrations (Spm, NADH, and NADPH, respectively) were 200 μM in all cases. a From Rodríguez et al. (2007). b From Frahry and Schopfer et al. (2007). Open in new tab PAO activity provides ·O2− and HO· radicals under saline stress When leaves of salt-treated plants were stained with NBT, they showed a strong decrease in precipitate intensity as compared with control leaves (Fig. 5), demonstrating a reduction of the ·O2− level due to the saline treatment. However, a certain amount of ·O2− was still observable in these conditions. The fact that NaCl substantially decreases NOX activity, the main source of apoplastic ·O2− and H2O2 in non-stressed maize plants (Schopfer et al., 2001; Rodríguez et al., 2007) as well as H2O2 production by NADH-dependent POX (Table 1), suggests the occurrence of some salt-tolerant mechanism for aROS production, alternative to POX and NOX. Interestingly, it has been proposed that AO activity is involved in the production of extracellular ·O2− and HO· radicals (Kawano et al., 2000a). In order to test whether this process takes place in vivo in the apoplast maize leaf under saline condition, ·O2− formation was determined by incubating salt-treated blade segments in XTT solution, in the presence or absence of SL-11061 (Fig. 6A). Our results showed that the addition of the PAO inhibitor resulted in a highly diminished ·O2− formation. These results (in addition to the fact that SL-11061 does not scavenge ·O2− radicals produced by NOX; see Table 1) support the idea that PAO activity indirectly produces most of the ·O2− in this zone under saline conditions. Similarly, when HO· was determined by the BZ method (Schopfer et al., 2001), PAO activity represented around 70% of the production of that free radical in the salt-treated segments (Fig. 6B). These results reinforced the hypothesis that, under saline conditions, PA oxidation by PAO would be the main source for aROS production in the elongation zone of maize leaf blades. Fig. 5. Open in new tabDownload slide ·O2− production in the whole leaf blade. ·O2− was detected by formazan precipitation. Control and salt-treated plants were incubated in the presence of 0.01% (w/v) NBT for 24 h. Plants were harvested and the third leaf was boiled in 80% (v/v) ethanol, mounted on a glass slide, and scanned. Abbreviations: c, unsalinized; s, salt-treated. Fig. 5. Open in new tabDownload slide ·O2− production in the whole leaf blade. ·O2− was detected by formazan precipitation. Control and salt-treated plants were incubated in the presence of 0.01% (w/v) NBT for 24 h. Plants were harvested and the third leaf was boiled in 80% (v/v) ethanol, mounted on a glass slide, and scanned. Abbreviations: c, unsalinized; s, salt-treated. Fig. 6. Open in new tabDownload slide In vivo PAO activity-derived ·O2− and HO· production under salinity. (A) ·O2− was detected by XTT according to Frahry and Schopfer (2001). Pools of eight salinized second blade segments were gently infiltrated and incubated for 7 h in the dark in 1 ml of aqueous solutions containing 0.5 mM XTT and 100 mM NaCl in the presence or absence of 100 μM SL-11061. The incubation medium (1 ml) was centrifuged at 10 000 g and the supernatant was subjected to measurement with a spectrophotometer at A470. Data were transformed into ·O2− molar extinction coefficient at 470 nm (2.16×104 M−1 cm−1). (B) HO· was detected by hydroxylation of BZ according to Schopfer et al. (2001). Pools of six salinized segments were gently infiltrated and incubated for 7 h in 1 ml of aqueous solutions containing 2.5 mM BZ and 100 mM NaCl plus or minus 100 μM SL-11061 at 30 °C in the dark. Fluorescence was determined in a spectrofluorometer (EX: 305 nm EM: 407 nm). Experiment was conducted twice, yielding similar results. Results are means ±SE (n=4). Asterisks indicate a difference from the control (P <0.05). Fig. 6. Open in new tabDownload slide In vivo PAO activity-derived ·O2− and HO· production under salinity. (A) ·O2− was detected by XTT according to Frahry and Schopfer (2001). Pools of eight salinized second blade segments were gently infiltrated and incubated for 7 h in the dark in 1 ml of aqueous solutions containing 0.5 mM XTT and 100 mM NaCl in the presence or absence of 100 μM SL-11061. The incubation medium (1 ml) was centrifuged at 10 000 g and the supernatant was subjected to measurement with a spectrophotometer at A470. Data were transformed into ·O2− molar extinction coefficient at 470 nm (2.16×104 M−1 cm−1). (B) HO· was detected by hydroxylation of BZ according to Schopfer et al. (2001). Pools of six salinized segments were gently infiltrated and incubated for 7 h in 1 ml of aqueous solutions containing 2.5 mM BZ and 100 mM NaCl plus or minus 100 μM SL-11061 at 30 °C in the dark. Fluorescence was determined in a spectrofluorometer (EX: 305 nm EM: 407 nm). Experiment was conducted twice, yielding similar results. Results are means ±SE (n=4). Asterisks indicate a difference from the control (P <0.05). In order to gain further insight into the involvement of higher polyamines catabolism by PAO on aROS generation, the effect was examined of adding different PAO modulators to the incubation buffer, on ·O2− production in the elongation zone of salt-treated plants, detected by XTT (Fig. 7). The addition of 100 or 200 μM Spm increased the in vivo ·O2− production, whereas treatment with SL-11061 (without Spm) or 1,8-DO, a commercial competitive PAO inhibitor (Ki=3×10−7 M; Cona et al., 2004), either separately or in combination with Spm showed the opposite effect (Fig. 7). As expected, Dap addition did not change the ·O2− levels. Similarly, ·O2− production in segments treated with diphenylene iodonium (DPI), reported as a NOX (Schopfer et al., 2001) and NADH-dependent POX inhibitor (Frahry and Schopfer, 1998) was similar to that of the control treatment, evidence that, under saline conditions, the enzymes mentioned were inhibited. Such in vivo inhibition was in line with that observed in the in vitro assay (Table 1) and in previous results concerning NOX (Rodríguez et al., 2007). Based on these results, it is concluded that PAO is the main contributor to apoplastic ·O2− production in salinized maize leaves. Fig. 7. Open in new tabDownload slide In vivo effect of PAO activity modulators on ·O2− production under salinity. Salinized second blade segments were used to detect ·O2− by XTT. Pools of eight segments were gently infiltrated and incubated for 7 h in the dark in 1 ml of aqueous solutions containing 0.5 mM XTT, 100 mM NaCl, and modulators of PAO activity. When indicated, the following concentrations were used: 200 μM SL-11061, 200 μM 1,8-DO, 100–800 μM Spm, 200 μM DPI, 200 μM Dap, 1 mM FZ, or 1 mM NC. Incubation medium (1 ml) was centrifuged at 10 000 g and the supernatant subjected to measurement with a spectrophotometer at A470. Data were transformed into ·O2− molar extinction coefficient 470 nm (2.16×104 M−1 cm−1). The experiment was performed twice, yielding similar results. Results are mean ±SE (n=8). Bars with the same letter are not significantly different (P <0.05). Fig. 7. Open in new tabDownload slide In vivo effect of PAO activity modulators on ·O2− production under salinity. Salinized second blade segments were used to detect ·O2− by XTT. Pools of eight segments were gently infiltrated and incubated for 7 h in the dark in 1 ml of aqueous solutions containing 0.5 mM XTT, 100 mM NaCl, and modulators of PAO activity. When indicated, the following concentrations were used: 200 μM SL-11061, 200 μM 1,8-DO, 100–800 μM Spm, 200 μM DPI, 200 μM Dap, 1 mM FZ, or 1 mM NC. Incubation medium (1 ml) was centrifuged at 10 000 g and the supernatant subjected to measurement with a spectrophotometer at A470. Data were transformed into ·O2− molar extinction coefficient 470 nm (2.16×104 M−1 cm−1). The experiment was performed twice, yielding similar results. Results are mean ±SE (n=8). Bars with the same letter are not significantly different (P <0.05). Possible source for the observed ·O2− and HO· H2O2 may be consumed to generate ·O2− and HO· through the Haber–Weiss reaction (Haber and Weiss, 1932) and POX activity in the Fenton-like reaction (Schopfer et al., 2001; Liszkay et al., 2004; Carol and Dolan, 2006). Therefore, the possibility that, under saline conditions, these free radicals originate in a reaction from the H2O2 produced by PAO (which remains active under salinity), through a chain reaction catalysed by Fe2+ or Cu+ (Fry, 1998; Kawano et al., 2000b) was tested. Salinized segments were treated with Spm plus the Fe2+-specific chelator FZ (Kosegarten et al., 1999) or the Cu+-specific chelator NC (Kunapuli and Vaidyanathan, 1983). Results demonstrated that chelators significantly decreased ·O2− production, indicating a probable involvement of a Fenton–Haber–Weiss-like reaction in this process. Effect of PAO and ROS modulators on segment elongation of salinized plants So far it has been shown that aPA oxidation is mostly responsible for the presence of certain aROS amounts in leaf blades of salinized maize plants. However, since the amount of aROS (H2O2 and H2O2-derived ·O2− and HO·) produced in leaf segments is much lower in the presence of salt, compared with that of the control without NaCl, the question remains as to whether the observed amounts of these aROS may still contribute to leaf elongation. To answer this question, the effect of diverse PAO and ROS modulators on segment elongation of salinized plants was tested (Fig. 8). Incubation with 200 μM Spm doubled segment elongation, whereas the addition of SL-11061 attenuated and 1,8-DO reversed the last effect. Interestingly, 800 μM Spm had no effect on segment elongation. In the absence of exogenous Spm addition, both PAO inhibitors diminished segment length, as compared with the control. Moreover, when 1,8-DO was added to the plant nutrient solution from the beginning of salt treatment, it produced reduced growth and Dap accumulation, as well as increased Spd and Spm contents of the entire maize leaf (Table 2), compared with plants not treated with the inhibitor. The addition of MnCl2, a ·O2− dismutating agent (Hernández et al., 2001) or BZ along with Spm, nullified the elongating effect of the polyamine on salinized segments. Finally, the incorporation of the specific Ca2+ chelating agent ethylene glycol bis (β-aminoethylether)-N, N, N′, N′-tetra-acetic acid (EGTA), reduced segments length, even in the presence of 200 μM Spm. Table 2. Effects of maize PAO inhibition on leaf growth and PA contents under salinity Control 1,8-DO Length (cm) 14.46±0.88 10.79±0.62* Polyamines(nmol g−1 FW)     Put 54.6±6.1 41.1±1.4*     Spd 134.9±4.6 209.3±37.6*     Spm 14.0±5.8 29.6±5.7*     Dap 238.2±0.6 116.5±4.8*     1,8-DO 0.0±0.0 13.3±1.92* Control 1,8-DO Length (cm) 14.46±0.88 10.79±0.62* Polyamines(nmol g−1 FW)     Put 54.6±6.1 41.1±1.4*     Spd 134.9±4.6 209.3±37.6*     Spm 14.0±5.8 29.6±5.7*     Dap 238.2±0.6 116.5±4.8*     1,8-DO 0.0±0.0 13.3±1.92* Plants were treated with nutrient solution plus 300 μM 1,8-DO for 14 d, from the beginning of salinization. Length of the expanding third leaf was measured and Dap, Put, Spd, Spd, and 1,8-DO levels of homogenized EZ segments determined by HPLC according to Jiménez-Bremont et al. (2007). Data are means ±SE (n=10 and n=4 for length and PA, respectively). Asterisks indicate significantly different from control (P <0.05). Open in new tab Table 2. Effects of maize PAO inhibition on leaf growth and PA contents under salinity Control 1,8-DO Length (cm) 14.46±0.88 10.79±0.62* Polyamines(nmol g−1 FW)     Put 54.6±6.1 41.1±1.4*     Spd 134.9±4.6 209.3±37.6*     Spm 14.0±5.8 29.6±5.7*     Dap 238.2±0.6 116.5±4.8*     1,8-DO 0.0±0.0 13.3±1.92* Control 1,8-DO Length (cm) 14.46±0.88 10.79±0.62* Polyamines(nmol g−1 FW)     Put 54.6±6.1 41.1±1.4*     Spd 134.9±4.6 209.3±37.6*     Spm 14.0±5.8 29.6±5.7*     Dap 238.2±0.6 116.5±4.8*     1,8-DO 0.0±0.0 13.3±1.92* Plants were treated with nutrient solution plus 300 μM 1,8-DO for 14 d, from the beginning of salinization. Length of the expanding third leaf was measured and Dap, Put, Spd, Spd, and 1,8-DO levels of homogenized EZ segments determined by HPLC according to Jiménez-Bremont et al. (2007). Data are means ±SE (n=10 and n=4 for length and PA, respectively). Asterisks indicate significantly different from control (P <0.05). Open in new tab Fig. 8. Open in new tabDownload slide Effects of PAO activity modulators, ROS scavengers, and a Ca2+ chelator on segment elongation. Second blade segments from salinized plants were incubated for 7 h in the dark with 100 mM NaCl. When indicated, the following concentrations were used: 200 μM SL-11061, 200 μM 1,8-DO, 50–800 μM Spm, 10 mM MnCl2, 5 mM BZ, and 10 mM EGTA. Segments were scanned before and after the incubation period and their length measured with an image processing software. The results are the percentage of length increase, with respect to the control during a 7 h incubation period. Absolute growth rate for control salinized second blade segments was 0.221±0.008 mm h−1. The experiment was conducted twice, yielding similar results. Results are means ±SE (n=20). Bars with the same letter are not significantly different (P <0.05). Fig. 8. Open in new tabDownload slide Effects of PAO activity modulators, ROS scavengers, and a Ca2+ chelator on segment elongation. Second blade segments from salinized plants were incubated for 7 h in the dark with 100 mM NaCl. When indicated, the following concentrations were used: 200 μM SL-11061, 200 μM 1,8-DO, 50–800 μM Spm, 10 mM MnCl2, 5 mM BZ, and 10 mM EGTA. Segments were scanned before and after the incubation period and their length measured with an image processing software. The results are the percentage of length increase, with respect to the control during a 7 h incubation period. Absolute growth rate for control salinized second blade segments was 0.221±0.008 mm h−1. The experiment was conducted twice, yielding similar results. Results are means ±SE (n=20). Bars with the same letter are not significantly different (P <0.05). Discussion Evidence has accumulated over recent decades demonstrating that polyamines play an important role in many plant developmental processes (Evans and Malmberg, 1989; Walden et al., 1997) and in plant responses to salinity and other abiotic stress conditions in diverse plant species (Krishnamurthy and Bhagwat, 1989; Galston and Sawhney, 1990; Aziz et al., 1998; Bouchereau et al., 1999; Simon-Sarkadi et al., 2002; Sanchez et al., 2005; Kusano et al., 2007). Several biotechnological approaches like overexpressing PA-synthesizing enzymes (Kumria and Rajam, 2002; Capell et al., 2004; Kasukabe et al., 2004; Wi et al., 2006) or antisense and mutant generation (Kasinathan and Wingler, 2004; Yamaguchi et al., 2006), allowed the generation of plants with increased and decreased stress tolerance, respectively. Recent studies using transgenic plants overexpressing or downregulating apoplastic polyamine oxidase, revealed the importance of the H2O2 derived from aPA catabolism in the induction of either salinity-induced tolerance or programmed cell death in tobacco (Moschou et al., 2008). The present work was focused on the possible involvement of the ROS produced from aPA oxidation in leaf growth processes of maize plants, grown under saline conditions. The first results showed that salt treatment reduced elongation in the region spanning 10–20 mm from the leaf ligule and, in parallel, it provoked a remarkable increment of higher apoplastic polyamines concentration in that region. This observation is in agreement with recent results showing that Spd, which is synthesized in the cytoplasm, is secreted into the apoplast upon salt treatment in tobacco (Moschou et al., 2008). In the present work, any other possibility was ruled out in order to confirm that salinity stimulates the passage of these substrates from the cytoplasm to the apoplast in maize plants. Similarly, the salt-induced decrease observed in Put levels, concomitant with the increase of total Spm levels of cell-free extracts in the segments, are consistent with the Spm accumulation described by other authors in salinized rice (Maiale et al., 2004), several vegetables (Zapata et al., 2004), Lotus glaber (Sannazzaro et al., 2007), and maize (Jiménez-Bremont et al., 2007). In vivo and in vitro measurements of H2O2 levels in the presence of saturating substrate conditions revealed the maximum feasible AO activity and showed, on the one hand, that PAO was the main enzyme contributing to the total PA oxidation level in maize leaves. Consequently, further studies were performed only on PAO activity, leaving CuAO activity aside. Biochemical, histochemical, and immunocytochemical studies allowed the localization of PAO, showing that it is specially abundant in the primary and secondary cell walls of xylem parenchyma, the endodermis, and epidermis of maize seedlings (Cona et al., 2006a). On the other hand, it was shown that salt treatment does not affect maximal PAO activity, suggesting that the enzyme is tolerant to this stress. Furthermore, the results obtained without exogenous substrate (that is to say, based on the actual polyamine cell content in the tissue) via the detection of the PAO product, Dap (Fig. 4A), demonstrated that salinity enhanced PAO activity. The last result consistently reflected the high Spd and Spm levels in that region (Fig. 2D–F) and suggested that the observed increase of inherent PAO activity under saline stress was a consequence of the rise in its substrate. These results are in line with in vitro results obtained by Smith (1977). Interestingly, the fact that the aDap amount was two and three orders higher than those of Spd and Spm, respectively, suggests that higher PA were actively oxidized to Dap and H2O2 in the apoplast, once they crossed the plasma membrane. Although out of the scope of the present work, the possibility that polyamine metabolism in the root (the first organ sensing salinity) behaves upon salt treatment in similar manner as the leaf blade is intriguing. As far as we know, the information regarding root PAO activity and salinity is limited to one report by Zhao et al. (2003), who reported that 0–200 mM NaCl increased Put, Spd, and PAO activity in the roots of barley seedlings. Unfortunately, this information was not discussed in terms of root growth or elongation. Evidence of reduced ·O2− amounts in salt-treated leaves by NBT staining (Fig. 5), revealed the occurrence of some mechanism for its production, alternative to that of NOX and NADH-dependent POX activities, which (unlike PAO) was strongly inhibited by salinity. The remarkable increase in in vivo ·O2− production by Spm addition, along with the substantially lowered in vivo ·O2− and HO· generation in the salt-treated segments by both PAO inhibitors or the Fe2+ and Cu+-specific chelators FZ and NC (Figs 6, 7), support the notion that ·O2− and HO· generation could occur from H2O2 production through PA oxidation and a further reaction catalysed by Fe2+ or Cu+, such as a Fenton–Haber–Weiss chain reaction (Kawano et al., 2000a, b). It is noteworthy that in vitro, cadaverine, putrescine, spermidine, and spermine do not scavenge superoxide radicals, but were found to be scavengers of hydroxyl radicals (Das and Misra, 2004) and unpublished results from our group have confirmed those results. However, such a ROS-scavenger effect was observed only when polyamines were used in concentrations of 0.5 mM or higher. As in the present work, polyamine concentration has been always much lower than that amount, we may discard any ROS-scavenging effect in our results. AO activity has formerly been related either to cell elongation in roots and hypocotyls of soybean seedlings (Delis et al., 2006) or to cell wall maturation in tobacco (Paschalidis and Roubelakis-Angelakis, 2005; Cona et al., 2006a). The purpose of this work was to evaluate the possible involvement of ROS production by PA oxidation in the leaf growth of maize plants grown under saline stress conditions. Our results suggest that tetramine oxidation contributes 25–30% of segment elongation under salinity (Fig. 8). Furthermore, when applied systemically along with NaCl, the PAO inhibitor 1,8-DO caused a 25% reduction in the elongation of whole leaves, compared with the control treatment without the inhibitor. Bearing in mind that the blade region analysed has 90% of the leaf elongation, it is deduced that, under a salt-stress situation, the minor contribution of PAO activity could still mean a significant yield improvement from an agronomical viewpoint. As a whole, these facts generate the expectation that biotechnological approaches like overexpressing enzymes responsible for PA biosynthesis or catabolism may be used to overcome reductions in the productivity of maize plants caused by salinity. In turn, the elimination of Spm-stimulated elongation by the specific Ca2+ chelating agent EGTA (Fig. 8), suggests that such elongation could be mediated by the activation of non-selective cation channels (NSCCs), through the HO· produced by Spm oxidation (a possibility that should be addressed in future research). This proposal is supported by the bulk of the evidence that has appeared during the last decade, which showed transient increases in cytosolic Ca2+ ([Ca2+]cyt) as a second messenger, suggesting that there are ROS/[Ca2+]cyt signalling pathways in several developmental processes. For example, guard cells and stomatal closure has been reported in Commelina communis and A. thaliana (McAinsh et al., 1996; Pei et al., 2000), as well as (2)-catechin-induced ROS production followed by ROS-induced Ca2+ increases in Centaurea diffusa and Arabidopsis thaliana roots (Bais et al., 2003) or the growth stimulation of A. thaliana roots (Foreman et al., 2003) and pollen tubes (Demidchik and Maathuis, 2007) by the aROS activation of Ca2+-permeable NSCCs that induce inward Ca2+ currents. Recently, it was shown that the lack of Spm in the Arabidopsis acl5/spms mutants caused hypersensitivity to NaCl, possibly due to impaired Ca2+-homeostasis (Yamaguchi et al., 2006) and that H2O2 generated by CuAO activates NSCCs in the abscisic acid-induced stomatal closure process in Vicia faba (An et al., 2008). ROS could also act on growth through a promotion of cell wall polysaccharide cleavage in vivo (Schopfer, 2001), such as that shown to operate in vitro (Miller, 1986; Fry, 1998; Schweikert et al., 2000). However, the action of ROS in the apoplast should be viewed as a delicate balance between cleavage and cross-linking activities (Cosgrove, 1999). An increase in PAO immunolabelling was observed inside secretory cytoplasmic organelles, suggesting the need for the intraprotoplasmic production of H2O2 for polymer cross-linking in the secretory pathway (Fry et al., 2000; Cona et al., 2003). Also, the balance between cleavage and cross-linking activities may be associated with a differential activity of cell wall peroxidases because different soluble peroxidase isozymes characterize the expanding and expanded regions in maize leaves (de Souza and MacAdam, 2001) and in Festuca arundinacea (MacAdam et al., 1992). The effectiveness of SL-11061 as inhibitor towards PA oxidation was formerly demonstrated in vivo, in an experiment using leaf blade segments in the presence of Spd (Maiale et al., 2008). In the present work, the inhibitory effect of DPI on NOX and NADH-dependent POX (Table 1) reported previously is also confirmed: 50 μM DPI inhibited NOX activity by 77% (Rodríguez et al., 2007) and the H2O2-producing activity by NADH-dependent POX by 94% (Frahry and Schopfer, 2001). In addition, strong NOX inhibition by DPI is in congruence with the Ki=5.6×10−6 M reported by O'Donnell et al. (1993). Reductions in ·O2− production and elongation of salinized segments treated with SL-11061 in vivo (Figs 7, 8), in addition to the fact that this polyamine analogue is efficient as a PAO but not as a NOX or NADH-dependent POX in vitro inhibitor (Table 1), demonstrated that PAO is not repressed by salinity. These facts also supported the use of this inhibitor to distinguish the PAO contribution to aROS production from that of the other two enzymes, under saline conditions. On the other hand, DPI treatment did not diminish either ·O2− production or elongation of salinized segments (Figs 7, 8), in agreement with in vitro observations (Table 1), showing that NOX and NADH-dependent POX activities are inhibited in vivo by salinity. Apoplastic Na+ concentration varies among and within plant species. Under salt treatment, apoplastic ion concentrations of 164 mM and 56 mM were reported in pea and spinach, respectively (Speer and Kaiser, 1991), whereas it approached 600 mM in salt-stressed rice plants (Flowers et al., 1991). Dissimilar apoplastic Na+ contents have been reported in salt-stressed maize leaves: 4–5 mM (Lohaus et al., 2000), 25 mM (Neves-Piestun and Bernstein, 2001), and 76 mM (own results). Compared with earlier reports, the higher apoplastic Na+ content registered in the present work may be ascribed to a more concentrated NaCl solution used for salinization (150 versus 100 or 80 mM) or to variations in other experimental conditions. Such a diversity of results on apoplastic Na+ contents highlights the importance of having assessed the actual apoplastic ion content in the salinized plant material under study in order to set a realistic experimental condition. Finally, variations in the effect of different Spm concentrations on ·O2− production and segment elongation (Figs 7, 8) gave evidence of a concentration-dependent Spm effect on ROS production and segment elongation. Unfortunately, previous reports describing changes in apoplastic PA levels have not measured cell elongation (Yoda et al., 2003; Angelini et al., 2008; Marina et al, 2008; Moschou et al., 2008). Taken together, our results demonstrated that, under saline stress, PAO might still provide the necessary H2O2 to generate ·O2− through an increased substrate availability and thus sustain leaf elongation. These results allowed us to propose the model depicted in Fig. 9. Thus, in the scenario where NOX is inhibited by non-lethal NaCl stress and the ROS produced by this enzyme is substantially reduced, oxidation by PAO of Spm and Spd accumulated in the apoplast of the EZ would result in an alternative source to generate ROS, partially counteracting the growth-inhibiting effect caused by salinity. Fig. 9. Open in new tabDownload slide Model of apoplastic ROS generation in a maize leaf blade grown under saline and non-saline conditions. The illustration integrates the models for probable apoplastic HO· production by POX (Schopfer et al., 2001) and PAO (through the Fenton–Haber–Weiss reaction, own results), higher PA catabolism by PAO (own results), and the activation of ROS-NSCC by Ca2+ (Foreman et al., 2003) in saline and non-saline conditions. SPDS: spermidine synthase; SPMS: spermine synthase. Fig. 9. Open in new tabDownload slide Model of apoplastic ROS generation in a maize leaf blade grown under saline and non-saline conditions. The illustration integrates the models for probable apoplastic HO· production by POX (Schopfer et al., 2001) and PAO (through the Fenton–Haber–Weiss reaction, own results), higher PA catabolism by PAO (own results), and the activation of ROS-NSCC by Ca2+ (Foreman et al., 2003) in saline and non-saline conditions. SPDS: spermidine synthase; SPMS: spermine synthase. Abbreviations Abbreviations [Ca2+]cyt cytosolic Ca2+ ·O2− superoxide radical 1,8-DO 1,8-diamino-octane 4-AAP μM 4-aminoantipyrine AES atomic emission spectrophotometry AOs amine oxidases aPA apoplastic extract aROS apoplastic ROS BZ sodium benzoate cPA cell extract CuAO copper-containing amine oxidase Dap 1,3-diaminopropane DCHBS 3,5-dichloro-2-hydroxybenzenesulphonic acid DPI diphenylene iodonium EGTA ethylene glycol bis (β-aminoethylether)-N, N, N′, N′-tetra-acetic acid EZ elongation zone FZ ferrozine H2O2 hydrogen peroxide HO· hydroxyl radical NBT nitro-blue tetrazolium NC neocuproine NOX NADPH oxidase NSCCs non-selective cation channels PA polyamines PAO polyamine oxidase PCA perchloric acid POX peroxidase Put putrescine ROS reactive oxygen species SL-11061 1,19-bis-(ethylamine)-5,10,15 triazanonadecane SOD superoxide dismutase Spd spermidine Spm spermine XTT Na, 3′-[1-[(phenylamino)-carbonyl]-3,4-tetrazolium](4-methoxy-6-nitro) benzenesulphonic acid hydrate The authors are grateful to Dr Edith Taleisnik (IFFIVE-CONICET, Argentina) and Dr Andrés Gárriz for their helpful advice, to Dr Benjamin Frydman for providing us with the SL-11061 inhibitor (SLIL Biomedical Corporation), and to Patricia Uchiya for technical assistance. 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Waters, Brian M.; Uauy, Cristobal; Dubcovsky, Jorge; Grusak, Michael A.

doi: 10.1093/jxb/erp257pmid: 19858116

Abstract The NAM-B1 gene is a NAC transcription factor that affects grain nutrient concentrations in wheat (Triticum aestivum). An RNAi line with reduced expression of NAM genes has lower grain protein, iron (Fe), and zinc (Zn) concentrations. To determine whether decreased remobilization, lower plant uptake, or decreased partitioning to grain are responsible for this phenotype, mineral dynamics were quantified in wheat tissues throughout grain development. Control and RNAi wheat were grown in potting mix and hydroponics. Mineral (Ca, Cu, Fe, K, Mg, Mn, P, S, and Zn) and nitrogen (N) contents of organs were determined at regular intervals to quantify the net remobilization from vegetative tissues and the accumulation of nutrients in grain. Total nutrient accumulation was similar between lines, but grain Fe, Zn, and N were at lower concentrations in the NAM knockdown line. In potting mix, net remobilization of N, Fe, and Zn from vegetative tissues was impaired in the RNAi line. In hydroponics with ample nutrients, net remobilization was not observed, but grain Fe and Zn contents and concentrations remained lower in the RNAi line. When Fe or Zn was withheld post-anthesis, both lines demonstrated remobilization. These results suggest that a major effect of the NAM genes is an increased efflux of nutrients from the vegetative tissues and a higher partitioning of nutrients to grain. Biofortification, grain protein content, iron, remobilization, senescence, zinc Introduction Wheat is a crop of major importance and together with other staple cereals supply the bulk of calories and nutrients in the diets of a large proportion of the world population (Cakmak, 2008). Cereals are inherently low in protein and mineral micronutrients such as Fe and Zn (White and Broadley, 2005, 2009; Cakmak, 2008; Newell-McGloughlin, 2008). A major focus of wheat breeders has been grain protein concentration as it affects bread- and pasta-making quality, but micronutrient improvement has received less attention. Approximately half of the world's population suffers from Fe and/or Zn deficiencies (Cakmak, 2008) and millions of children suffer from protein-energy malnutrition (de Onis et al., 1993). As such, the improvement of nutritional quality of wheat could benefit the nutritional status of millions of people. A common agronomic practice to increase grain protein concentration is the use of N fertilization. However, this practice is expensive and excess fertilizer run-off is a potential environmental contaminant (Masclaux-Daubresse et al., 2008). A substantial percentage of the N in wheat grain is supplied by amino acids remobilized from vegetative tissue (Barneix, 2007; Gregersen et al., 2008; Masclaux-Daubresse et al., 2008). Much of this N content is derived from proteins that are disassembled and recycled during the leaf senescence stage of development (Hopkins et al., 2007). Likewise, Fe and Zn have been shown to be remobilized from vegetative tissues in several plants (Hocking and Pate, 1977; Hocking, 1994; Miller et al., 1994; Drossopoulos et al., 1996; Waters and Grusak, 2008), although the specific sources are unknown. Zinc fertilization has been a successful strategy to improve wheat grain Zn concentration (Cakmak, 2008), and improvement in the partitioning or remobilization of Zn to grain could make fertilization efforts more efficient. Wheat grain with higher Zn concentration has been demonstrated to produce more vigorous crops (Cakmak, 2008; Yilmaz et al., 1998). Thus, breeding or transgenic approaches that result in plants with increased partitioning of minerals to grain could be useful for both nutritional biofortification and reduced fertilizer application. Chromosome 6B from wild emmer wheat (Triticum turgidum ssp. dicoccoides) was identified as a potential source of genetic variation for grain protein (Joppa and Cantrell, 1990), Zn, and Fe concentration (Cakmak et al., 2004). A quantitative trait locus (QTL) for grain protein concentration was mapped on chromosome arm 6BS (Joppa et al., 1997) and later mapped as a single Mendelian locus, Gpc-B1 (Olmos et al., 2003). In near-isogenic lines of this locus, increased grain protein was associated with the increased remobilization of amino acids from the flag leaf (Kade et al., 2005), higher grain Fe and Zn concentrations (Cakmak et al., 2004; Distelfeld et al., 2007), and accelerated leaf yellowing, indicating accelerated senescence (Uauy et al., 2006a). A NAC transcription factor, NAM-B1, was identified as the causal gene for Gpc-B1 by positional cloning (Uauy et al., 2006b). Other members of the NAC family are known to regulate developmental processes (Aida and Tasaka, 2006; Olsen et al., 2005), including leaf senescence (Guo and Gan, 2006). In transgenic wheat NAM RNA interference (RNAi) lines in which NAM-B1 and its homeologous genes had decreased expression, leaf yellowing was delayed, and grain protein, Fe, and Zn concentrations were greatly decreased (Uauy et al., 2006b). These results, together with higher N, Fe, and Zn concentrations in RNAi line flag leaves at maturity, suggested a role for NAM-B1 homeologues in the remobilization of N compounds, Fe, and Zn. However, without taking organ mass, nutrient concentrations at prior time points, and total nutrient accumulation of other organs into account, this model could not be confirmed. In addition, the body of literature does not contain sufficient data regarding sources of grain minerals (other than N) to support the idea that remobilization alone could account for the differences observed. Because a whole-plant partitioning profile has not been undertaken in plants differing in NAM-B1 expression, it is currently unclear whether this gene directly affects remobilization (which is defined here as the net loss of stored mineral content from an organ over time), alters partitioning of nutrients within the plant, alters total plant uptake of these nutrients, or influences a combination of these processes. The current study uses multiple time point sampling of an expanded profile of mineral concentrations and contents (which takes organ mass into account) of all shoot organs in NAM knockdown and control lines. This sampling allows the quantification of N and mineral remobilization as contributors to final grain protein and mineral content, and provides a better understanding of the physiological effects of the NAM genes. In addition, experimental treatments have been included to test the remobilization capacity of Fe and Zn in these lines, and radiolabelled Zn was used to quantify the short-term translocation to grain. The information presented here will inform future genomic and systems level studies designed to understand genes and processes that can be targeted to increase grain mineral concentrations for the biofortification of foods. Materials and methods Plant growth in potting mix The lines used in this study were the NAM RNAi lines designed to reduce expression of all NAM family members (event L19-54) and its non-transgenic control (Uauy et al., 2006b). The transformed line is Bobwhite, a semi-dwarf, hard, white, spring, common wheat (T. aestivum) variety. Wheat seeds were imbibed at 4 °C for 3 d and allowed to germinate in darkness at room temperature for 4 d. Seedlings were planted in commercial potting mix (MetroMix 300; Sun Gro Horticulture, Bellevue, WA, USA) and vermiculite at a 2:1 ratio in 17 cm diameter, 17 cm tall pots, three plants per pot, and placed in a growth chamber (16 h photoperiod; 350 μE m−2 s−1 of photosynthetically active radiation at the top of the pots, 22.5/17.5 °C day/night, relative humidity set at 50%). Pots were placed in trays with two pots of each line per tray. Plants were watered as needed by sub-irrigation (usually twice per week) with a nutrient solution (2.0 l per tray) of the following composition: 1.2 mM KNO3, 0.8 mM Ca(NO3)2, 0.8 mM NH4NO3, 0.3 mM KH2PO4, and 0.2 mM MgSO4. Plants were sampled at anthesis of the first emerged head (d0), and at 14, 28, 35, 42, and 56 d after anthesis (DAA), with an additional harvest at 70 DAA for the RNAi line. At each sampling, the number of tillers was noted, and plants were cut with a scalpel into the following parts: heads, peduncles, stems, lower leaves, and flag leaves. For each plant, organs from all tillers were pooled and a total of 4–6 plants (replicates) from two separate pots were analysed per time point. Tissues were dried for 48 h in a drying oven at 60 °C, and dry weights were obtained. After drying, heads were separated into grain, rachis, and florets, grains were counted, and these parts were weighed. Plant growth in hydroponics Wheat seeds were imbibed as described above. Seedlings were planted in plastic cups with plastic beads for support, then placed in lids over 4.5 l containers, 10 plants per pot, in a growth chamber with the settings as described above. Plants were maintained in a complete nutrient solution of the following composition: 3.0 mM KNO3, 1.0 mM Ca(NO3)2, 0.5 mM KH2PO4, 0.5 mM MgSO4, 0.75 mM K2SO4, 0.1 mM K2SiO3, 25 μM CaCl2, 25 μM H3BO3, 0.5 μM MnSO4, 0.5 μM ZnSO4, 0.5 μM CuSO4, 0.5 μM H2MoO4, 0.1 μM NiSO4, and 10 μM Fe(III)-HEDTA (N-hydroxyethylethylene-diaminetriacetic acid; Sigma Chemical Co., St Louis, MO, USA). Solutions were buffered at pH 5.8 with 2.0 mM MES (2-[N-morpholino] ethane sulphonic acid, monohydrate; Research Organics, Cleveland, OH, USA), and changed twice weekly. For Fe or Zn deficiency treatments (0 Fe and 0 Zn, respectively), ZnSO4 or Fe(III)-HEDTA were omitted from the solution after anthesis. Plants were sampled at anthesis of the first emerged head (d0), and at 42 DAA for the Fe deficiency and control treatments (+Fe+Zn). Since 0 Zn plants matured more rapidly than control or 0 Fe plants, plants were sampled at 35 DAA for Zn deficiency and control treatments. At each sampling, the number of tillers was noted, and plants were cut with a scalpel into the following parts: heads, peduncles, stems, lower leaves, and flag leaves. Usually, all tillers from a plant were collected, although the occasional late-emerging tiller was discarded. Thus, 2–5 tillers from a minimum of two plants per time point were collected. Organs from each tiller were analysed separately, then average values were calculated. Tissues were dried for 48 h in a drying oven at 60 °C, and dry weights were obtained. After drying, grain was removed from heads, and weighed separately. Elemental analysis For mineral analysis of potting mix-grown plants, above-ground organs were dissected and organs from all tillers of each plant were pooled as described above. All tissues except florets were ground in a stainless-steel coffee mill. Duplicate subsamples of approximately 250 mg were weighed into glass tubes, and digested in nitric:perchloric acid (4:1 v/v) for 1 h at 100 °C, then gradually to 200 °C until the sample was taken to dryness. Samples were then resuspended in 15 ml 2% nitric acid. All acids were trace metal grade (Fisher Scientific, Pittsburgh, PA, USA) and water was filtered through a MilliQ system (Millipore, Billerica, MA, USA) to 18 MΩ resistivity. Mineral concentrations were determined by ICP-OES (CIROS ICP model FCE12; Spectro, Kleve, Germany). The mineral content of the tissues was calculated by multiplying tissue DW by each mineral concentration. For mineral analysis of hydroponically grown plants, a different digestion procedure was used (due to a laboratory decision to curtail the use of perchloric acid). Whole organs were weighed into glass tubes, and digested in 2.0 ml nitric acid overnight, then at 125 °C for 1.5 h. 1.5 ml 30% H2O2 was added and samples were digested for 1 h. A second 1.5 ml volume of H2O2 was added and samples were digested for 1 h. The temperature was then increased to 200 °C and samples were evaporated to dryness. Residues were dissolved in 15 ml 2% nitric acid. Mineral concentrations were determined by ICP-OES. The mineral content of the tissues was calculated by multiplying tissue DW by each mineral concentration. For N analysis, tissue samples were dried to constant weight and ground to a fine powder using a ball mill. The samples were analysed for N concentration by a continuous-flow mass spectrometer (Europa Scientific, Cambridge, UK) at the University of California, Davis Stable Isotope Facility. Radiotracer studies Wheat plants were grown in hydroponics as described above. The dates of anthesis were noted, and at mid-grain fill (20–25 DAA), plants were moved from a complete nutrient solution to a complete solution spiked with 65Zn (Brookhaven National Laboratory, Upton, NY, USA) at 1 μCi l−1 for continuous labelling experiments, and at 4 μCi l−1 for pulse labelling experiments. All labellings were initiated between 3–4 h into the photoperiod. Plants were not removed from the growth chamber during the labelling period. For continuous labelling, the plants remained in the labelling solution for up to 24 h. For pulse-labelling, the plants were removed from the labelling solution after 3 h and rinsed for 10 min in complete, unlabelled nutrient solution, then placed in fresh complete, unlabelled nutrient solution. At 12 h (at or near end of photoperiod) or 24 h (3–4 h into following photoperiod) after the commencement of labelling, shoots were excised and cut into lower leaves, stems, flag leaf, peduncle, and heads. Heads were oven-dried for 4–12 h, then the grains were removed. All tissues were quantified for 65Zn by gamma counting. Statistics Analyses of variance were performed using the SAS Version 9.1 program (SAS Institute, Cary, NC, USA). The general linear model (PROC GLM) was used to assess the effect of the reduced NAM transcript levels in the RNAi lines as compared to the isogenic controls. Data were transformed when necessary using logarithmic and power transformations in order to meet the assumptions of the model. For comparisons over growth periods, each time point was analysed separately using orthogonal contrasts. Differences in mineral content and concentration between tissues at different time points were analysed using unpaired t tests. Values after the ‘±’ sign are standard errors of the mean throughout the text. Results Potting mix experiment Control and RNAi plants grew similarly in terms of appearance and total plant size (see Supplementary Fig. 1 at JXB online). This similarity was also true for individual plant organs, although some tissues differed at some time points. Total grain weight, on a per head basis, was similar between the two lines. Weight of individual kernels was nearly identical and reached maximum values by 35–42 DAA, although total grain dry weight (DW) continued to increase as a result of increased seed numbers at later time points. Across sampling points, RNAi lines had a higher number of grains per head, although these differences were significant only at 14 DAA and 35 DAA (P <0.03). It was observed that, as described previously by Uauy et al. (2006b), the most notable difference between the two lines was delayed leaf yellowing of the RNAi line. The complete data set of mineral contents for the potting mix-grown plants is presented in Supplementary Table S1 at JXB online. At anthesis, Fe and the contents of most other minerals (in μg) were similar in the vegetative organs of both lines (see Supplementary Table S1 at JXB online), suggesting that NAM genes had no effect on the content of these minerals to this point in development. Total Zn content was slightly lower in the RNAi line at anthesis (P=0.04). At grain maturity (56 DAA), the total shoot contents (vegetative tissues plus grain) of Fe, Zn, and N were similar in both lines (P >0.65), indicating that total uptake and accumulation of each mineral was not significantly affected by the NAM genes. Total vegetative Fe content (the sum of all non-grain organs) decreased between anthesis and maturity (i.e. exhibited net remobilization) in the control line (14.7%, P=0.36; Fig. 1; Table 1). Although the decrease in content between anthesis and maturity was not significant, the RNAi line total vegetative Fe content did not decrease, but rather increased significantly (P <0.02). Total vegetative Zn decreased only in the control line (60.9%, P <0.001). Comparing the quantity of mineral remobilized from all vegetative tissues to the quantity of mineral in the grain pool at maturity (56 DAA), the net remobilized Fe and Zn could account for between 13.0% of grain Fe and 42.6% of total grain Zn content in the control line, assuming that all of each mineral demonstrating net remobilization was translocated to the grain (Table 2). In the control line, Fe content decreased over time in lower and flag leaves, stems, peduncle, and rachis, indicating net remobilization. Contrary to this, Fe remained constant or accumulated over time in all tissues of the RNAi line (Table 1). This was especially marked in the peduncle of RNAi plants, which accumulated 286% of the initial Fe content. Zinc content decreased significantly in all vegetative tissues of the control line (P <0.01; Fig. 1; Table 1) In the RNAi line, Zn was remobilized from both the flag leaf and lower leaves, but the percentage change was lower than the control (Table 1). Zinc decreased in the RNAi line until 35 DAA in stem, peduncle, and head tissues, after which the content increased (see Supplementary Table S1 at JXB online). Table 1. Per cent change in Fe and Zn content from anthesis to maturity, and per cent change in N content from 35 DAA to 56 DAA Experiment Treatment Nutrient Flag leaf Lower leaves Peduncle Stem Florets Rachis Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA Fe −32.8 6.8 −7.4 33.1 −47.8 285.7 −8.0 77.3 32.6 53.4 −33.3 28.6 −14.7 38.9 Potting mix 56 DAA Zn −71.7 −50.9 −61.3 −24.4 −61.3 82.4 −61.4 −2.9 −63.9 43.9 −60.0 54.5 −60.9 1.8 Potting mix 56 DAA N −32.0 −5.0 −22.0 6.0 −26.0 17.0 −20.0 52.0 −6.0 20.0 nda nd −21.0 17.0 Experiment Treatment Nutrient Flag leaf Lower leaves Peduncle Stem Florets Rachis Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA Fe −32.8 6.8 −7.4 33.1 −47.8 285.7 −8.0 77.3 32.6 53.4 −33.3 28.6 −14.7 38.9 Potting mix 56 DAA Zn −71.7 −50.9 −61.3 −24.4 −61.3 82.4 −61.4 −2.9 −63.9 43.9 −60.0 54.5 −60.9 1.8 Potting mix 56 DAA N −32.0 −5.0 −22.0 6.0 −26.0 17.0 −20.0 52.0 −6.0 20.0 nda nd −21.0 17.0 Experiment Treatment Nutrient Flag Leaf Lower Leaves Peduncle Stem Head Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Hydroponic 35 DAA +Zn Zn −48.3 −14.0 15.5 20.8 0.0 −15.9 −34.9 −27.8 153.8 51.4 18.3 6.8 Hydroponic 35 DAA 0Zn Zn −80.6 −60.3 −69.9 −86.8 −92.8 −86.8 −90.1 −83.9 −49.2 −50.5 −74.8 −69.6 Hydroponic 42 DAA +Fe Fe −54.0 −16.0 −18.0 −16.0 −37.0 13.0 −20.0 25.0 111.0 48.0 −7.9 −3.7 Hydroponic 42 DAA 0Fe Fe −74.0 −21.0 −57.0 −36.0 −77.0 −40.0 −56.0 −15.0 −59.0 −55.0 −65.6 −40.2 Experiment Treatment Nutrient Flag Leaf Lower Leaves Peduncle Stem Head Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Hydroponic 35 DAA +Zn Zn −48.3 −14.0 15.5 20.8 0.0 −15.9 −34.9 −27.8 153.8 51.4 18.3 6.8 Hydroponic 35 DAA 0Zn Zn −80.6 −60.3 −69.9 −86.8 −92.8 −86.8 −90.1 −83.9 −49.2 −50.5 −74.8 −69.6 Hydroponic 42 DAA +Fe Fe −54.0 −16.0 −18.0 −16.0 −37.0 13.0 −20.0 25.0 111.0 48.0 −7.9 −3.7 Hydroponic 42 DAA 0Fe Fe −74.0 −21.0 −57.0 −36.0 −77.0 −40.0 −56.0 −15.0 −59.0 −55.0 −65.6 −40.2 a nd, Not determined. Open in new tab Table 1. Per cent change in Fe and Zn content from anthesis to maturity, and per cent change in N content from 35 DAA to 56 DAA Experiment Treatment Nutrient Flag leaf Lower leaves Peduncle Stem Florets Rachis Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA Fe −32.8 6.8 −7.4 33.1 −47.8 285.7 −8.0 77.3 32.6 53.4 −33.3 28.6 −14.7 38.9 Potting mix 56 DAA Zn −71.7 −50.9 −61.3 −24.4 −61.3 82.4 −61.4 −2.9 −63.9 43.9 −60.0 54.5 −60.9 1.8 Potting mix 56 DAA N −32.0 −5.0 −22.0 6.0 −26.0 17.0 −20.0 52.0 −6.0 20.0 nda nd −21.0 17.0 Experiment Treatment Nutrient Flag leaf Lower leaves Peduncle Stem Florets Rachis Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA Fe −32.8 6.8 −7.4 33.1 −47.8 285.7 −8.0 77.3 32.6 53.4 −33.3 28.6 −14.7 38.9 Potting mix 56 DAA Zn −71.7 −50.9 −61.3 −24.4 −61.3 82.4 −61.4 −2.9 −63.9 43.9 −60.0 54.5 −60.9 1.8 Potting mix 56 DAA N −32.0 −5.0 −22.0 6.0 −26.0 17.0 −20.0 52.0 −6.0 20.0 nda nd −21.0 17.0 Experiment Treatment Nutrient Flag Leaf Lower Leaves Peduncle Stem Head Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Hydroponic 35 DAA +Zn Zn −48.3 −14.0 15.5 20.8 0.0 −15.9 −34.9 −27.8 153.8 51.4 18.3 6.8 Hydroponic 35 DAA 0Zn Zn −80.6 −60.3 −69.9 −86.8 −92.8 −86.8 −90.1 −83.9 −49.2 −50.5 −74.8 −69.6 Hydroponic 42 DAA +Fe Fe −54.0 −16.0 −18.0 −16.0 −37.0 13.0 −20.0 25.0 111.0 48.0 −7.9 −3.7 Hydroponic 42 DAA 0Fe Fe −74.0 −21.0 −57.0 −36.0 −77.0 −40.0 −56.0 −15.0 −59.0 −55.0 −65.6 −40.2 Experiment Treatment Nutrient Flag Leaf Lower Leaves Peduncle Stem Head Total vegetative Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Hydroponic 35 DAA +Zn Zn −48.3 −14.0 15.5 20.8 0.0 −15.9 −34.9 −27.8 153.8 51.4 18.3 6.8 Hydroponic 35 DAA 0Zn Zn −80.6 −60.3 −69.9 −86.8 −92.8 −86.8 −90.1 −83.9 −49.2 −50.5 −74.8 −69.6 Hydroponic 42 DAA +Fe Fe −54.0 −16.0 −18.0 −16.0 −37.0 13.0 −20.0 25.0 111.0 48.0 −7.9 −3.7 Hydroponic 42 DAA 0Fe Fe −74.0 −21.0 −57.0 −36.0 −77.0 −40.0 −56.0 −15.0 −59.0 −55.0 −65.6 −40.2 a nd, Not determined. Open in new tab Table 2. Total net remobilization from vegetative tissues as percentage of seed mineral content Experiment Treatment Cu Fe K Mn P S Zn Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA 31.6 – 13.0 – – – – – 10.0* – – – 42.6* – Hydroponic 35 DAA +Zn 34.2*b 66.4* – – – 14.7 – – – – – – – – Hydroponic 35 DAA 0 Zn –a 7.8 – – – – – – – – – – 180.3* 185.5* Hydroponic 42 DAA +Fe 15.6* 64.9* 7.4 5.5 – – – – 53.5* – – – – – Hydroponic 42 DAA 0Fe 5.0 – 165.2* 163.2* 17.8 0.5 74.9* − 51.1* − 33.5* – – – Experiment Treatment Cu Fe K Mn P S Zn Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA 31.6 – 13.0 – – – – – 10.0* – – – 42.6* – Hydroponic 35 DAA +Zn 34.2*b 66.4* – – – 14.7 – – – – – – – – Hydroponic 35 DAA 0 Zn –a 7.8 – – – – – – – – – – 180.3* 185.5* Hydroponic 42 DAA +Fe 15.6* 64.9* 7.4 5.5 – – – – 53.5* – – – – – Hydroponic 42 DAA 0Fe 5.0 – 165.2* 163.2* 17.8 0.5 74.9* − 51.1* − 33.5* – – – a No net remobilization occurred. b Significant between anthesis and maturity at P <0.05. Open in new tab Table 2. Total net remobilization from vegetative tissues as percentage of seed mineral content Experiment Treatment Cu Fe K Mn P S Zn Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA 31.6 – 13.0 – – – – – 10.0* – – – 42.6* – Hydroponic 35 DAA +Zn 34.2*b 66.4* – – – 14.7 – – – – – – – – Hydroponic 35 DAA 0 Zn –a 7.8 – – – – – – – – – – 180.3* 185.5* Hydroponic 42 DAA +Fe 15.6* 64.9* 7.4 5.5 – – – – 53.5* – – – – – Hydroponic 42 DAA 0Fe 5.0 – 165.2* 163.2* 17.8 0.5 74.9* − 51.1* − 33.5* – – – Experiment Treatment Cu Fe K Mn P S Zn Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Control RNAi Potting mix 56 DAA 31.6 – 13.0 – – – – – 10.0* – – – 42.6* – Hydroponic 35 DAA +Zn 34.2*b 66.4* – – – 14.7 – – – – – – – – Hydroponic 35 DAA 0 Zn –a 7.8 – – – – – – – – – – 180.3* 185.5* Hydroponic 42 DAA +Fe 15.6* 64.9* 7.4 5.5 – – – – 53.5* – – – – – Hydroponic 42 DAA 0Fe 5.0 – 165.2* 163.2* 17.8 0.5 74.9* − 51.1* − 33.5* – – – a No net remobilization occurred. b Significant between anthesis and maturity at P <0.05. Open in new tab Fig. 1. Open in new tabDownload slide Stacked bar graph representing mineral contents in individual vegetative tissues and grain in potting mix-grown control and RNAi wheat at anthesis and at maturity (56 DAA). (A) Fe content, (B) Zn content. Values ±SE. Asterisk represents statistical difference (P <0.05) between time points. Total N was quantified in selected tissues and time points for the control and RNAi line. At anthesis and grain maturity, total N contents were similar between both lines (P=0.92 and P=0.68), suggesting that reduced NAM expression did not alter total N uptake and accumulation. Throughout grain development, total and individual grain N content were significantly lower in the RNAi line than in the control line (Fig. 2; P <0.05). As with Zn, RNAi plants remobilized N from flag leaves and lower leaves over the time-course, but to a lesser extent than in the control plants (29% RNAi versus 45% control flag leaf, 8.4% RNAi versus 33.7% control lower leaves). The greatest differences in flag leaf N remobilization occurred between 35 DAA and 56 DAA (Table 1). Comparing N content of all shoot vegetative organs (except the rachis, which was used entirely for mineral analysis) between these time points (Fig. 3), total shoot N (vegetative tissues plus grain) increased by similar quantities between the RNAi (8.0 mg) and control lines (7.5 mg). However, grain of the control line gained 14 mg of N, while grain of the RNAi line increased only by 2.7 mg. Vegetative tissues of control plants showed a net remobilization of 6.6 mg of N, while those of the RNAi line did not show a net remobilization of N, but rather N increased by 5.3 mg. From anthesis to maturity, N was remobilized from the control stem (18%) and peduncle (48%), while N content increased in these tissues of the RNAi line (+29% peduncle, +119% stem). This resulted in significant differences between control and RNAi lines for both grain N and vegetative N at maturity (56 DAA) (P <0.01). Fig. 2. Open in new tabDownload slide Nitrogen content of flag leaves and grain in control and RNAi plants over the growth period. (A) Flag leaf N content, (B) total grain N content (per tiller), (C) N content of individual grains. Values ±SE. Asterisk denotes statistical significance (P ≤0.05). Fig. 3. Open in new tabDownload slide Stacked bar graph representing nitrogen content in individual vegetative tissues and grain in control and RNAi wheat at 35 DAA and at 56 DAA (maturity). Values ±SE. Asterisk represents statistical difference (P <0.05) between time points. Hydroponic experiments To characterize further the mineral remobilization in the RNAi lines under different levels of Fe and Zn, plants were grown using hydroponic conditions, either in a complete nutrient solution for the duration of the experiment, or in an Fe- or Zn-deficient solution from anthesis onwards. Withholding Fe and Zn forced plants to rely solely on stored Fe and Zn to supply the grain. This treatment was used to assess potential net remobilization from the vegetative tissue to the grain while preventing uptake and xylem translocation of these minerals during grain fill, although some root-associated Fe and Zn may have supplied a finite quantity of residual mineral content. If the RNAi line was capable of remobilizing minerals, a net loss of Fe or Zn content would be detected in vegetative tissues between anthesis and maturity. The full data set is presented as Supplementary Table S2 available at JXB online. Leaf yellowing was delayed in the RNAi line in hydroponics, similar to potting mix-grown plants. The control and RNAi lines had similar Fe and Zn vegetative contents at anthesis (Fig. 4; P >0.75). For total shoots (less grain), neither line exhibited significant net remobilization of Fe or Zn when grown on complete nutrient solution, but both remobilized significant quantities of these minerals when grown on deficient nutrient solutions (P <0.02). Iron-deprived RNAi plants had significantly less net remobilization, with a decrease of 40.2% of vegetative Fe content as compared to 65.6% for the control line (Table 1; P=0.01). Zinc-deprived RNAi plants showed a net remobilization of 69.6% of Zn content compared to 74.8% for the control line, but the difference between lines was not significant (Table 1; P=0.66). The quantities of Fe and Zn remobilized were more than enough to account for grain mineral content (Table 2), although total Fe or Zn content in the grain for each line was significantly lower than when these minerals were supplied continuously (Fig. 4; see Supplementary Table S2 at JXB online; P <0.03). Some shoot Fe and Zn may have been translocated to roots to maintain root growth. Roots of both lines grown in complete nutrient solution did not decrease in Fe or Zn content during grain fill (see Supplementary Table S3 at JXB online). In Fe- and Zn-withholding treatments, root mineral content and concentration decreased substantially in both lines. Roots of all lines and treatments continued to grow during grain fill, and were larger at grain maturity than at anthesis, making calculation of remobilization by subtraction (as done with shoot organs) impractical. Fig. 4. Open in new tabDownload slide Stacked bar graph representing (A) Zn content and (B) Fe content in total vegetative tissues and grain in hydroponic grown control and RNAi lines at anthesis and maturity [35 DAA for (A); 42 DAA for (B)]. +Zn, Zn was supplied continuously; +Fe, Fe was supplied continuously; 0 Zn, Zn was omitted from nutrient solution from anthesis onward; 0 Fe, Fe was omitted from nutrient solution from anthesis onward. Values ±SE. Asterisk to the left of bar respresents statistical difference (P <0.05) from anthesis. Asterisk to the right of bar represents statistical difference (P <0.05) between lines at that time point. Despite the lack of significant net Fe and Zn remobilization from the total vegetative tissues of plants grown in complete nutrient solution, the control line exhibited substantial Fe and Zn net remobilization from some vegetative organs (Table 1). The control line exhibited a 54% and 48.3% decrease in flag leaf Fe and Zn content, respectively, between anthesis and maturity (35 DAA for Zn, 42 DAA for Fe; Table 2). For both minerals, the RNAi line remobilized significantly less (P <0.05) Fe (16%) and Zn (14%) from the flag leaves. This indicates that the NAM genes influence net remobilization from flag leaves even in the complete nutrient solution. The other minerals tended to accumulate or remain constant in flag leaves between anthesis and 35 DAA, and the effect of the NAM genes was not readily apparent (see Supplementary Table S2 at JXB online). An exception was Cu, which was also remobilized in both RNAi and control plants (Table 2; Supplementary Table S2 at JXB online). These data suggest that under these experimental conditions, the effect of NAM genes on mineral remobilization primarily affects remobilizable metal micronutrients, i.e. Fe, Zn, and Cu. Grain mineral accumulation Over the time-course of the potting mix experiment, grain Ca and Mg concentrations (μg g−1) were similar between lines at most time points, while K was higher in the RNAi line (Fig. 5). Copper, Fe, Mn, P, S, and Zn were at lower concentrations in the RNAi line at most time points. In the hydroponics experiments, the RNAi line produced more, although slightly smaller grains per head than the control line, resulting in significantly higher total grain mass per tiller (see Supplementary Fig. S2 at JXB online). Across treatments, the total grain mineral content per tiller was higher in the RNAi line than in the control line for Ca and K, and similar for Mg, Mn, P, and S (with the exception of 35 DAA 0 Zn; see Supplementary Table S2 at JXB online). This similar total grain mineral accumulation spread across a higher total grain weight in the RNAi line resulted in lower concentrations of Mg, Mn, P, and S in this line, relative to the control (Fig. 6). Copper, Fe and Zn concentrations were also lower, but by a greater percentage than the other minerals. In terms of total grain accumulation (content), Fe was lower in the RNAi line relative to the control in complete solution at 35 DAA and 42 DAA (38%, P=0.05), and in the Fe withheld experiment (36%, P <0.01). Grain Zn content in the RNAi line was also significantly lower than the control in complete solution at 35 DAA and 42 DAA (P <0.05), but not in the Zn deficiency treatment (Fig. 6; see Supplementary Table S2 at JXB online). Fig. 5. Open in new tabDownload slide Grain mineral concentrations over the grain development period in potting mix-grown wheat. DAA, days after anthesis. Values ±SE. Asterisk denotes statistical significance (P ≤0.05). Fig. 6. Open in new tabDownload slide Grain mineral concentrations of mature control and RNAi lines grown in hydroponics with complete nutrient solution (+Zn, 35 DAA, +Fe, 42 DAA), or nutrient solution lacking Zn or Fe from anthesis onward (0 Zn, 35 DAA; 0 Fe, 42 DAA). Values ±SE. Unless denoted by NS (not significant), control and RNAi values were statistically different (P ≤0.05). Radiotracer experiment To determine the effect of NAM genes on the short-term translocation of newly absorbed Zn to the grains, a radioactive tracer strategy was used. At mid-grain fill, plants were transferred from a complete nutrient solution to a complete nutrient solution spiked with 65Zn. After 12 h or 24 h, radioactivity was detected in all shoot organs. In three of the four experiments, the proportion of total shoot 65Zn that was partitioned into grain was significantly higher in the control line than in the RNAi line (Fig. 7). This was true regardless of whether the plants were continuously labelled or pulse-labelled. Fig. 7. Open in new tabDownload slide Short-term partitioning of newly taken up Zn to grain at mid-grain fill. Percentage of total shoot 65Zn counts in grain at each sampling for control and RNAi lines grown in complete nutrient solution spiked with 65Zn. 12 h and 24 h represent 12 h and 24 h sampling time points. Continuous indicates continuously labelled experiment and pulse the pulse-label experiment. Values ±SE. Asterisk denotes statistical significance (P ≤0.05). Discussion Remobilization: an operational definition Much of the N imported into wheat grain is derived from protein in vegetative tissues that is degraded to amino acids and recycled by translocation to the grain, i.e. remobilization (Barneix, 2007; Gregersen et al., 2008; Masclaux-Daubresse et al., 2008). It is often assumed that certain minerals supplied to seeds also come from remobilized sources, but, unlike N, specific sources of stored or recycled minerals are unknown, and the few studies that have quantified the contributions of minerals remobilized from vegetative tissues to seeds have reached differing conclusions (Hocking, 1994; Miller et al., 1994; Garnett and Graham, 2005; Peng and Li, 2005; Waters and Grusak, 2008). In this work, net remobilization is defined as the loss of stored mineral content over time from one organ, and subsequent accumulation of that mineral content into another organ. Because net change in mineral content over time is a function of influx and efflux of nutrients, net remobilization will be detected only when efflux exceeds influx. Thus, substantial quantities of a given mineral could pass through an organ without a detectable change in content, resulting in no detectable net remobilization. Likewise, minerals could be remobilized from one subcellular compartment (or organ) while accumulating in another compartment or in the apoplastic spaces (or in another organ) without a change in total content or net remobilization. Since all shoot tissues have been collected and analysed in these experiments, it is possible to assess mineral partitioning to various tissues over time. If the grain mineral pool were to increase while the shoot mineral pool remained constant, then the quantity of mineral translocated to the grain must have passed through the shoot tissues, and would be equal to the quantity entering the shoot during that time period. When comparing lines, if translocation of mineral (from remobilization or pass through) to the grain is inhibited, the decreased flux will be detected as a relative increase in vegetative mineral content and less of an increase in grain mineral content. Although this discussion will focus on Fe, Zn, and N, other minerals were quantified in order to determine whether the effects of the NAM genes were general in nature, or if certain minerals were disproportionately affected. Quantifying tissue DW and multiple minerals also demonstrated that remobilization did not occur for all minerals, and that observed changes in organ content were not secondary effects of changes in growth or organ mass. Effect of NAM genes on remobilization under different mineral availabilities The NAM genes are members of the NAC transcription factor family and were previously shown to affect grain Fe, Zn, and N content in a dosage-dependent manner. Construction of the RNAi line used here and the resultant alterations in NAM transcript levels have been described previously (Uauy et al., 2006b). As transcription factors, NAM proteins are predicted to regulate genes that encode for proteins that carry out physiological processes for nutrient remobilization and/or translocation to grain. The RNAi and control lines used in our work only differ in their relative NAM gene expression, and are otherwise isogenic. Therefore, differences in Fe, Zn, and N dynamics between the control and transgenic lines can be assigned to direct or downstream effects of these genes. Our results indicate that the extent of net remobilization is dependent on availability of mineral inputs and thus will probably be highly dependent on environmental conditions in field-grown plants. In complete hydroponic nutrient solution growth conditions, no significant net remobilization of Fe or Zn was observed in either line. Despite this, grain Fe and Zn contents and concentrations were substantially higher than those from plants grown in potting mix, where remobilization of both Fe and Zn was observed in control lines. When hydroponic plants were deprived of Fe or Zn inputs post-anthesis, net remobilization occurred in both the control and RNAi lines, from shoot tissues (Table 2) and also from roots (see Supplementary Table S3 at JXB online). Both lines remobilized more than enough of these minerals to account for the entire grain content, although the Fe and Zn quantities accumulated in the grain were substantially lower than in plants on the complete solution treatment. These results suggest that while remobilization and partitioning of Fe and Zn to grain is impaired in the NAM knockdown line, this is not due to a complete inhibition of remobilization, as the RNAi line is capable of remobilizing minerals under nutrient-limiting conditions. They also suggest that when Fe and Zn are readily available to the roots, and are adequately absorbed into the plant, this source supersedes the need for remobilization from the leaves. In the absence of sufficient Fe and Zn from the soil, the plant can obtain these minerals from the storage forms present in both shoot and root vegetative tissues. These results are consistent with those obtained when P was withheld from wheat plants during grain development (Peng and Li, 2005). Remobilization: quantification and putative mechanisms In potting mix growth conditions, net remobilization of Fe and Zn from the control line was observed, but diminished or no net remobilization in the RNAi line. Total accumulation of plant Fe and Zn was similar, but partitioning of these minerals to grain was substantially lower in the RNAi line. In this same experiment, vegetative N content decreased in the control line over time, indicating net remobilization, while there was an increase in N in the RNAi line. Between 35 DAA and 56 DAA, net remobilized N could account for 46% of the increase in grain N in the control line, but accounted for none of the N in the grain of the RNAi line. These results, in combination with the 65Zn experiment that demonstrated decreased short-term translocation of Zn to grain, indicate that the translocation of certain minerals and N to grain is impaired in the NAM knockdown line. A combination of decreased efflux and sustained influx of minerals into vegetative tissues could account for the lower net remobilization exhibited in the RNAi lines, and could also account for the lower percentage of total Fe, Zn, and N partitioned to grain. Target genes of the wheat NAM transcription factors have not been identified, and the molecular mechanism by which the NAM proteins affect translocation to grain (net remobilization plus pass through) is currently unknown. A microarray study of senescence in Arabidopsis leaves revealed a large number of up-regulated transporter proteins, including OPTs, YSLs, and ZIPs (Van der Graaff et al., 2006). It is possible that NAM proteins regulate similar transporter genes in wheat and that these genes are needed for effective Fe and Zn remobilization. Other possible explanations include indirect effects on phloem loading for efflux of Fe, Zn and N (as amino acids) from leaves; or an effect on the rate or timing of disassembly of the internal sources of these elements. The latter hypothesis is supported by the observation that lines with a functional copy of the NAM-B1 gene had higher soluble protein and amino acids concentration in the flag leaves than near isogenic lines with a non-functional NAM-B1 gene (Kade et al., 2005). The reduced expression of NAM genes and the accompanying delay of normal vegetative development, i.e. senescence, may result in a disruption of the normal source and sink tissue relationship. Delayed senescence and the accompanying degradation of proteins may result in a situation where the substrates for transporters (amino acids or minerals) are decreased or not present, or are only present (or available) later in the grain-filling period and thus are less efficiently translocated out of source tissues. Hundreds to thousands of proteins are estimated to interact with Zn ions as structural or catalytic components or as substrates (Broadley et al., 2007), thus substantial quantities of Zn could be released during protein degradation. Similarly, Fe from the degradation of chloroplast proteins could be released during leaf senescence. Delayed degradation of chloroplasts containing these proteins, as suggested by the delay in leaf yellowing in the RNAi line, may explain why remobilization of Fe was inhibited proportionally more than the remobilization of Zn. Since the grain of the RNAi plants grew normally (based on appearance and DW gain), the movement of water and photoassimilates did not seem to be impaired. This suggests an inhibition of translocation processes more specific to Fe, Zn, and N rather than a general inhibition of phloem transport. Practical implications The average grain Zn concentration of potting mix-grown control plants was similar to that of field-grown wheat (Rengel et al., 1999; White and Broadley, 2005; Morgounov et al., 2007). However, grain of control line wheat grown in complete hydroponic culture had a Zn concentration approximately five times higher (195 μg g−1 versus 38 μg g−1), which parallels the improvements in Zn grain concentrations made via Zn fertilization (Cakmak, 2008). The grain Fe concentration of the hydroponic control line was approximately twice that of potting mix-grown plants (99 μg g−1 versus 44.7 μg g−1). In the RNAi line also, Fe and Zn concentrations in grain from plants grown on complete hydroponic solution were significantly higher than in grain from potting mix-grown plants (Fe, 51 μg g−1 hydroponic versus 29.9 μg g−1 potting mix; Zn, 87 μg g−1 hydroponic versus 29 μg g−1 potting mix). These results suggest that wheat grain is already capable of accumulating several-fold higher Fe and Zn concentrations than are usually obtained in field situations. Because the RNAi line had lower partitioning of Fe and Zn to grain under both high and low availability, grain concentrations of these nutrients can possibly be increased by improvements in the efficiency of translocation. Indeed, overexpression of an Arabidopsis Zn transporter in barley resulted in increased seed Zn concentration (Ramesh et al., 2004). However, constitutive overexpression of a Zn transporter in rice resulted in the aberrant distribution of Zn within the plant (Ishimaru et al., 2007). Overexpression of transporters may need to be targeted spatially and temporally to result in the desired increases of nutrients in the target tissue. The transgenic line also showed reduced translocation of N. It is estimated that grain protein in the control line was 19.7%, while protein in the RNAi line was one-third lower, at 13.0%. These values are higher than normally observed in field situations, possibly as a result of the continuous supply of N to the plants. While decreased expression of the NAM genes negatively affects the accumulation of Fe, Zn and protein in the grains, increasing the transcript levels of the NAM genes above the levels normally found in current commercial varieties can result in increased protein. The B genome copy of the NAM1 gene (NAM-B1) is non-functional or deleted in modern bread wheat (Uauy et al., 2006b), and introgression of a functional copy from wild wheat can significantly increase Fe, Zn, and N grain concentration in certain genotype–environment conditions (Uauy et al., 2006a), and can also result in a significant increase in total N content (grain N concentration×grain yield) (Brevis and Dubcovsky, 2009). However, accelerated senescence in several isogenic lines containing a functional NAM-B1 allele resulted in reduced grain-filling periods and reduced kernel weights. Therefore, the best genotype–environment combinations must be determined in the breeding process to deploy NAM-B1 cultivars effectively. Conclusions and future directions Use of the NAM knockdown line for comparative physiology has allowed us to understand further the movement of minerals through the plant and eventually to seeds. The results suggest that NAM genes affect remobilization and/or pass through of Fe, Zn, and N from vegetative tissues to grain. The transgenic lines analysed here provide a valuable entry point for deciphering specific genes and processes involved in nutrient movement to seeds. Knowledge of genes operating downstream of the NAM genes could provide new targets to engineer a more efficient efflux and/or remobilization of Fe, Zn, and N from source tissues at the proper developmental stages. In addition to the improvements in wheat nutritional value, the improvement of mineral partitioning to grain would be an environmentally and economically beneficial improvement, because less fertilizer might be required to produce grain of similar N and mineral concentration. Projects in this direction are currently in progress in our laboratories. This work was funded in part by funds from USDA-ARS under Agreement No 58-6250-6-003 and from the Harvest Plus Project under Agreement No 58-6250-4-F029 to MAG and by funds from USDA-CSREES grant 2008-35318-18654 to JD. The authors would like to thank Adrian J Bituin and Francine J Paraiso for laboratory assistance. The contents of this publication do not necessarily reflect the views or policies of the US Department of Agriculture, nor does mention of trade names, commercial products, or organizations imply endorsement by the US Government. References Aida M , Tasaka M . 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Effect of seed zinc content on grain yield and zinc concentration of wheat grown in zinc-deficient calcareous soils , Journal of Plant Nutrition , 1998 , vol. 21 (pg. 2257 - 2264 ) Google Scholar Crossref Search ADS WorldCat Author notes † Present address: Department of Crop Genetics, John Innes Centre, Colney Lane, Norwich NR4 7UH, UK. © 2009 The Author(s). This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.5/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details) © 2009 The Author(s).

Thomas, R. G.; Hay, M. J. M.

doi: 10.1093/jxb/erp258pmid: 19717528

Abstract Axillary buds within a plant shoot system are known to differ in their ability to respond to treatments favouring their development. This ability is referred to as their outgrowth potential. Using two species of prostrate nodally-rooting herbs, dicotyledonous Trifolium repens and monocotyledonous Tradescantia fluminensis, grown throughout in a strictly vegetative state, this study tested two hypotheses. Hypothesis 1: that each axillary bud exhibits an outgrowth potential that is directly related to the growth rate of its parent apical bud, and Hypothesis 2: that the growth rate attained by an axillary bud depends upon both its outgrowth potential and the local supply of stimulatory root-derived signal (NRS) available to it. Activation levels (growth rates) of apical buds were varied by differential exposure to nodal roots and the outgrowth responses of axillary buds recently emerged from them were then measured under standardized conditions of NRS supply. Hypothesis 1 was shown to be correct for both species. Hypothesis 2, tested only in T. repens, was supported by results showing that an axillary bud's outgrowth potential and the NRS supply to it each independently influenced its growth rate, there being no significant interaction between the two. These results emphasize the significant role the physiological state/activity of apical buds has on the outgrowth potential of axillary buds formed within them. The fact that similar relationships were observed on axillary buds on stems of differing developmental maturity and branching hierarchy, and in two taxonomically diverse species, suggests they might be widespread among morphologically similar species. Axillary bud outgrowth, branch development, bud activation, bud outgrowth potential, nodal roots, prostrate clonal herbs, root signals, Tradescantia fluminensis, Trifolium repens Introduction Studies with nodally-rooting, prostrate-stemmed perennial species indicate that their shoot branching is regulated predominantly by fluctuations in stimulatory signal(s) transported acropetally from roots (Thomas et al., 2002, 2003a, b; Thomas and Hay, 2004, 2007, 2008a, b). This is in marked contrast to much of the reported work on the regulation of branching in the erect-stemmed annual species of Arabidopsis thaliana, Pisum sativum, and Petunia hybrida, where genetic and physiological evidence points to the dominance of common inhibitory influences from roots and apical buds (see reviews by Leyser, 2005; McSteen and Leyser, 2005; Beveridge, 2006; Dun et al., 2006; Ongaro and Leyser, 2008; and Johnson et al., 2006; Simons et al., 2007, Gomez-Roldan et al., 2008; Umehara et al., 2008). In these model systems, it has been demonstrated clearly that there is a network of shoot and root feedback and interacting signals that collectively operate to regulate branching (Beveridge, 2006; Dun et al., 2006; Aguilar-Martínez et al., 2007; Simons et al., 2007; Ongaro and Leyser, 2008; Ferguson and Beveridge, 2009) and, in the case of Petunia, evidence has been presented of a stimulatory influence of roots that is expressed in the presence of the inhibitory influence (Napoli, 1996; Simons et al., 2007). Thus it appears that in these model species root signals have both stimulatory and inhibitory components although the net effect is usually inhibitory. Alternatively, in prostrate-stemmed species, the balance appears to favour the stimulatory signal with the reduction in axillary bud development at nodes distanced from the basal root resulting from a decrease in effectiveness of the stimulus. Thomas and Hay (2008a) suggested that decreased effectiveness might arise as a consequence of a decrease in the stimulatory signal, an increase in supply of the inhibitor, decreasing sensitivity of the axillary buds to the stimulatory signal with distance from the root system, or a combination of these factors. In view of this, the stimulatory influence transported from nodal roots in the prostrate-stemmed plants of this study is likely to be the net result of both stimulatory and inhibitory influences and is referred to henceforth, for convenience, as the net root stimulus (NRS). At present the nature of the stimulatory signal is unknown. As foliar spray application of mineral nutrient to sub-optimally supplied plants restored the growth of the shoot organs but not branching (Hay et al., 2003), nutrient supply by nodal roots is unlikely to be directly involved. However, in these prostrate-stemmed species, xylem transport of root-derived cytokinin cannot be ruled out as the possible stimulatory signal, with its level being regulated by polar auxin transport to roots (Li and Bangerth, 1999; Bangerth et al., 2000) although there is now strong evidence from erect-stemmed species indicating that the cytokinin promoting axillary bud outgrowth is locally biosynthesized in the nodal region of the stem (Tanaka et al., 2006; Ferguson and Beveridge, 2009) with auxin regulating both its biosynthesis and its degradation (Shimizu-Sato et al., 2009). It was shown earlier (Thomas and Hay, 2007) in Trifolium repens that the rate of outgrowth of an axillary bud (its activation level) in response to a stimulus from a nodal root is cumulative and is maintained after excision of the nodal root. Using this species as being representative of the group of nodally-rooting, prostrate-stemmed species (Thomas and Hay, 2004, 2008b), Thomas and Hay (2008a) attempted to develop a predictive model of the rate of growth of axillary buds based on the level of activation each bud attained from its exposure to NRS at the time of its emergence from its parent apical bud. While this model was able to explain the decline in growth rate of successively produced axillary buds on a primary stem as its apical bud grows away from its basal root system, it failed to explain the rapid progressive decline in secondary branch development on successively formed lateral branches. They therefore suggested that there is a need to take into account an additional factor and were led to hypothesize that the ability of an emerging axillary bud to respond to a given level of local NRS might be, at least in part, directly influenced by the activation level (growth rate) of its parent apical bud during its early development within it. A similar association between the outgrowth of axillary buds and the developmental stage and physiological activity of the shoot terminal meristem has been suggested to occur in Pisum (Gould et al., 1987), Nicotiana (McDaniel and Hsu, 1976), and Arabidopsis (Grbic and Bleecker, 2000). This intrinsic ability of an axillary bud to respond to NRS supply is conceptually similar to that previously described as its ‘outgrowth potential’ (Napoli et al., 1999) or ‘growth potential’ (Husain and Linck, 1966; Gould et al., 1987). In the present paper this ability is referred to as ‘outgrowth potential’. To assess the validity of the concept that an axillary bud's potential to respond to NRS supply is related to the level of activity of the apical bud in which it formed, experiments were undertaken to test two specific hypotheses. Hypothesis 1: that each axillary bud exhibits an outgrowth potential that is directly related to the growth rate of its parent apical bud. Hypothesis 2: that the rate of growth attained by an axillary bud depends upon the combination of its outgrowth potential and the local supply of NRS available to it at the time of its emergence from its parent apical bud. To achieve this, it was necessary to design experiments in which the activation levels (growth rates) of apical buds were varied while holding all other variables constant. The steps involved in the experiments were thus: firstly, the manipulation of shoot and root systems to provide apical buds with a range of different activation levels. Secondly, the prevention of possible variation in the inhibitory influence of parent apical buds on the outgrowth of axillary buds (test buds) newly emerged from them, by excising the apical bud tissues immediately distal to the test buds. Thirdly, ensuring, by appropriate excision procedures, that test buds from parent apical buds with different activation levels were exposed to a similar supply of NRS from the source root systems. Fourthly, in Trifolium, the manipulation of nodal roots to provide two different levels of NRS supply to test buds formed by parent apical buds with differing activation levels. To broaden the relevance of the findings from this study, experiments were carried out on two phylogenetically widely distanced species; namely, the dicotyledonous Trifolium repens and a monocotyledonous species, Tradescantia fluminensis, with a similar growth habit and apical bud structure (Denne, 1966). Materials and methods Plant material Trifolium repens L. (white clover): experimental plants were derived from a greenhouse-grown stock clone of a single genotype of T. repens selected from a Spanish ecotype collection (AgResearch Accession number C1067) previously described (see Thomas et al., 2003b; Thomas and Hay, 2007, 2008a). Tradescantia fluminensis Vell. (wandering jew): This monocotyledonous species (family Commelinaceae) is a prostrate perennial clonal herb the stems of which, as in T. repens, produce roots at nodes wherever they contact moist ground. Cuttings from a single genotype growing in a nearby garden were grown in a heated greenhouse to produce stock plants from which the experimental plants were derived. These remained vegetative throughout the experiment. Culture of experimental plants of both species Plants were grown from stem tip cuttings planted in a commercially obtained potting mix (Thomas et al., 2002) in 1.35 l planter bags. After about 3 weeks, the two or three basalmost branches formed by this time were trimmed off each plant to leave a single primary stem growing away from its basal root system. All lateral branches that grew out subsequently from this primary stem were retained. The oldest phytomer on the primary stem that retained a branch at its node was termed phytomer 1 (P1) and later-formed ones termed P2, P3, etc. Outgrowth of nodal roots was prevented by growing shoot systems out over a dry plastic mesh. Throughout the investigation, plants were grown in a heated greenhouse in natural photoperiods at average maximum/minimum temperatures of 25/12 °C. Where required, outgrowth of a nodal root on a primary stem was stimulated by firmly pinning the youngest newly emerged node to a mound of moist potting mix contained in a 200 ml plastic pot. Experimental design Trifolium experiment: outgrowth potential of axillary buds on branches: Plants were grown for a 10-week pretreatment phase (Fig. 1a) during which apical buds on successively formed branches attained successively lower activation levels (Thomas et al., 2002). They were then given one of four core treatments (Fig. 1b, A–D) or two additional treatments (Fig. 1b, E, F) and responses of axillary buds that had formed within branch apical buds with different activation levels were measured over a 5-week period. Fig. 1. Open in new tabDownload slide Experimental design and experimental procedures followed for the Trifolium experiment: (a) state of the plants during the pretreatment phase, at the time of rooting of the distal node at P10 (i) or P14 (iii) and at the time of treatment application (ii) and (iv), and (b) state of the plants of each treatment immediately after the imposition of treatments showing the oldest, middle, and youngest (X, Y, and Z, respectively) test axillary buds (TB) and the axillary branches/buds (Ax) that were excised (each one indicated by a small x). Arrowheads show the positions of apical buds, black dots represent nodes bearing axillary buds, and open circles depict phytomers where nodal root outgrowth was stimulated. Broken lines in (b) indicate the positions of actively growing lateral branches but not their relative lengths. Short double lines indicate points of tissue excision. Pretreatment: Thirty-six plants were grown from shoot tip cuttings until 5 May 2004, by which time ten phytomers had emerged from the apical buds of their primary stems (Fig. 1a(i)). At this stage the NRS supply to phytomer 10 (P10) from the basal root system was sufficient for continued growth of the primary stem apical bud but inadequate for strong outgrowth of newly emerging axillary buds. To produce branch apical buds of differing activation levels, nodal root outgrowth was stimulated at the youngest node on the primary stem (at P10) on 30 plants [Fig. 1a(ii)], and, 19 d later, at P14 on a further six plants [Fig. 1a(iv)]. Development of these nodal root systems stimulated axillary bud activation and subsequent outgrowth of branches at the phytomers at and distal to them, but the apical bud on the branch at the rooted node became more highly activated (grew faster) than those on successively more distal branches at non-rooted nodes. All plants were then grown on for 45 d until the start of treatments on 19 June 2004 when the branch at P14 had three emerged phytomers. At this time the apical bud on the P14 branch was either weakly activated when the nodal root was distanced four phytomers from it at P10 [Fig. 1a(ii)] or more strongly activated when the root was at P14 [Fig.1a(iv)]. Branches at P1–P9 were similar in all plants and were retained throughout the experiment. Core treatments: The core treatments (Fig. 1b, A–D) were set up to compare outgrowth rates of three test axillary buds (X, Y, and Z) on the P10 and P14 branches in response to NRS (Fig. 1b). These were the youngest three newly emerged buds immediately proximal to their parent branch apical bud, and therefore of similar developmental age in each treatment. They were also similar as to their phytomeric distance (number of phytomers) from the root at P10 (Fig. 1b), thereby ensuring that they were subjected to a similar supply of root-derived stimulus (NRS) during their development (Thomas and Hay, 2008a) up to the time that treatments were applied. To avoid possible variation in competition for NRS from the branches and buds intercalated between the test buds and the node at P10, all these intercalated branches and buds were excised at the start of core treatments A–D. At the same time, all primary stem tissue was excised distal to the point of attachment of the branches at P10 (treatments A, C) or P14 (treatments B, D), and to avoid the possibility of differences between inhibitory influences from faster and slower growing apical buds on branches at P10 and P14, respectively, these apical buds were excised immediately distal to the test buds. These excisions resulted in all the NRS available at P10 being channelled solely to the test buds in each treatment. Some leaflets were removed as necessary to even up the total area of leaves subtending the test buds. To assess the influence of NRS supply on test bud responses, the P10 nodal root was either repotted into a 1.0 l container and retained, in treatments A and B, or excised at the start of treatments, in treatments C and D (Fig. 1b). Thus in treatments A and B, distal buds continued to receive NRS from both the basal root system and the nodal root (high NRS), or, in treatments C and D, they became dependent solely on NRS from the basal root (low NRS). Differences brought about solely by differences in the supply of NRS are apparent by comparison of treatment A with C and treatment B with D. Differences between axillary bud outgrowth responses on branches at P10 and P14 resulting solely from differences between the activation levels of their parent apical buds (outgrowth potential) are discernible by comparing treatment A with B and treatment C with D. Treatment E was included to verify that the activation levels of the apical buds of the P10 and P14 branches did differ as predicted, by measuring their growth rates at the start of the treatment period. This treatment was identical with treatments C and D with regard to excision of the nodal root at P10, but differed in that no excisions of shoot material were made (Fig. 1b). Because P10 branches formed earlier on the primary stems than those at P14, one inevitable consequence of the core experimental design (treatments A–D) was that test buds differed in their positions on the branches bearing them, being at the sixth, seventh, and eighth nodes that formed on the larger P10 branch compared with the first, second, and third on the smaller, younger, branch at P14 (see Fig. 1b). Treatment F, in which six plants were stimulated to root at P14 [Fig. 1a(iv)] on 24 May 2004, 19 d after nodal root formation at P10 in treatments A–E, was therefore included. This treatment, in which the nodal root at P14 was excised after 26 d, was otherwise directly comparable to treatment D in which the nodal root at P10 was excised after 45 d. It was essential as a means of distinguishing whether differences found between the outgrowth responses of test buds on branches at P10 and P14 resulted simply from their positions on the branches rather than from differences in outgrowth potentials. As it was known that the presence of a nodal root for 3–4 plastochrons is sufficient to activate fully the axillary bud at a rooted phytomer (Thomas and Hay, 2007), it was predicted that the presence of nodal roots for 26 d in treatment F would raise the activation level of the apical buds of the branches at P14 so that the outgrowth rates of their test buds would be significantly increased relative to those in treatment D and would approach those of the test buds in treatment C. Such a response in bud outgrowth rates on P14 branches in treatment F would indicate that their outgrowth potential, rather than their position on the branches, was modifying their outgrowth rate. The effects of all six treatments were monitored weekly for 5 weeks. In treatments A–D and F, assessments were made of leaf emergence from, and stem length of, branches developing from the test buds. In treatment E, measurements were made of rates of stem elongation of the branches at P10 and P14 and of leaf emergence from their apical buds. Tradescantia experiment: outgrowth potential of axillary buds on the primary stem: This experiment was aimed at establishing the relationship between the outgrowth potential of axillary buds on the primary stem and the activation level (growth rate) of their parent apical buds at the time the axillary buds were formed within them. To achieve this, the activation levels of primary stem apical buds were varied by differential durations of exposure to the stimulatory influence of nodal roots [Fig. 2b(i), Phase 1] (Thomas and Hay, 2007, 2008a) and the outgrowth rates of axillary buds that had newly emerged from them were then measured in response to excision of their parent apical buds under common levels of NRS supply from the basal root systems during Phase 2 [Fig. 2b(ii)]. Fig. 2. Open in new tabDownload slide Experimental design and experimental procedures followed for the Tradescantia experiment: (a) state of the plants at the end of the pretreatment phase, at the time of rooting of the distal node at P19, and (b) state of the plants when treatments were applied (i) during Phase 1 where the treatments of nodal root duration at P19 were implemented by excising the nodal root after 8, 12, 18, or 24 d of growth and then growing the plants on until Day 63, and (ii) during Phase 2 where on Day 64 the apical bud of the primary stem was excised from three of the six plants of each rooting treatment and all plants then grown on for a further 23 d during which the test axillary buds were monitored for leaf emergence and stem elongation. TB, test axillary buds; X, Y, and Z, oldest, middle, and youngest axillary buds, respectively. Arrowheads show the positions of apical buds, black dots represent nodes bearing axillary buds, and broken lines indicate the positions (but not relative lengths) of actively growing lateral branches. Short double lines indicate points of tissue excision. Pretreatment: Thirty plants were established on 14 August 2006, such that each had a non-rooted primary stem extending away from a basal root system. By the end of the pretreatment period, the primary stems bore about 12 lateral branches, the axillary buds at their six youngest phytomers had failed to grow out into branches, and their 19th non-rooted phytomers were newly emerged from their apical buds (Fig. 2a). Treatments: Treatments were started on 1 October 2006 by pinning the node at P19 onto moist potting mix to stimulate nodal root development on 24 of the 30 plants [Fig. 2b(i)], the remaining six being controls in which nodal root development was prevented (treatment 0D). Phase 1 treatments differed as to the length of time nodal roots were permitted to develop by excising them after 8, 12, 18, or 24 d in treatments 8D, 12D, 18D, and 24D, respectively. All plants were then grown on for a further 9 weeks from the onset of stimulation of root development, by which time differential activation of primary stem apical buds was apparent. A further set of treatments (Phase 2) was given on 5 December 2006 (Day 64), by excising the apical bud on the primary stem of three plants in each treatment and leaving the remaining three plants intact to act as controls [Fig. 2b(ii)]. All plants were then grown on for a further 3 weeks, until 28 December 2006, to allow the three youngest axillary buds immediately proximal to the excised apical bud on the primary stem of the decapitated plants to respond to the treatments. In all treatments, all previously formed branches proximal to the test axillary buds were retained for the duration of the experiment. The effects of the duration of rooting treatments during Phase 1were assessed by measuring the rates of elongation of the primary stem and of leaf emergence from its apical bud during the period from nodal root stimulation through to the time of excision of the apical bud (5 December 2006). Elongation of the three test axillary buds (X, Y, and Z) on the primary stem of each decapitated plant and the equivalent buds on the control plants was then monitored for 3 weeks during Phase 2 (5–28 December 2006). Lastly, at the end of the experiment each shoot was divided into a basal and an apical region by severing the primary stem immediately proximal to P19. These portions were then oven-dried at 60 °C for 3 d before weighing. Analysis of data A single mean value for leaf emergence and stem elongation of the test buds for each treatment was calculated from the independently assessed values of each of the test buds (X, Y, and Z) in each plant within each treatment. In the Trifolium experiment, the number of nodes and the branch stem length data at Day 35 in Fig. 3 were analysed using the planned contrast option of analysis of variance in GenStat statistical software (Payne et al., 2007). The effects of root retention and branch position and their interaction were all compared at the between-plant level of variability. Means and least significant differences (LSD5%) for comparing them were obtained from the relevant interaction tables. Fig. 3. Open in new tabDownload slide Test bud outgrowth in the Trifolium experiment: (a) the cumulative number of leaves emerged per week, and (b) cumulative elongation of the bud stems per week for the mean of the three test axillary buds in treatments A–D and F. Values are presented for each of the 5 weeks of the assessment period. The vertical bar represents the LSD5% for treatment differences in the cumulative 35 d values. In the Tradescantia experiment, the slopes of fitted linear regression lines of each plant in each treatment were tested by ANOVA (Payne et al., 2007) for the significance of treatment (duration of nodal root presence) on elongation of the primary stem during Phase 1 (Fig. 4). Appropriate diagnostic checks were performed after model fitting. ANOVA was used to test for the significance of the treatment on the stem length of test buds at 23 d (Fig. 5). The simple linear regression analyses reported in Fig. 6 were performed using the Microsoft Excel software package. Fig. 4. Open in new tabDownload slide The cumulative lengths (mm) of the primary stems distal to the nodal root at P19 at intervals over the 9 week Phase 1, growth period in the Tradescantia experiment. Nodal roots were present at P19 at the beginning of the Phase 1 period for 0 d (0D), 8 d (8D), 12 d (12D), 18 d (18D), or 24 d (24D). Bars represent ±SEM. Vertical arrows show times of nodal root excision after 8, 12, 18, and 24 d. Fig. 5. Open in new tabDownload slide Cumulative elongation of bud stems per week over 23 d for the mean of the three test axillary buds in each treatment of the Tradescantia experiment. Values are presented for each of the three assessments over this period. The vertical bar represents the LSD5% for treatment differences in the cumulative 23 d values. Fig. 6. Open in new tabDownload slide The relationship in Tradescantia fluminensis between the activation levels of parent apical buds (measured as the rates of elongation of primary stem tissues produced by them) and those of the test axillary buds (measured as the stem elongation rates of the branches developed from them). The rate of primary stem elongation was calculated over the last week of Phase 1, whereas that for the test axillary buds was calculated over the second and third weeks of Phase 2. Results Trifolium experiment: bud outgrowth responses on branches (a) Hypothesis 1: outgrowth potential of an axillary bud is directly related to the growth rate (activation level) of its parent apical bud Growth rates of the apical buds of experimental branches at P10 and P14: The results of treatment E (Fig. 1b) established that growth rates of the apical buds that were retained on the branches at P10 and P14 were, indeed, different. Leaf emergence rates from these buds, and the rates of elongation of the internodes emerging from them, were both substantially greater on the P10 than on the P14 branch (Table 1). Furthermore, their growth rates were constant; the rates of both leaf emergence and stem elongation being similar in weeks 1 and 3. Table 1. Mean rates of leaf emergence on, and stem elongation of, the branches at P10 and P14 of treatment E during the first and third weeks of the assessment period of the Trifolium experiment Branch position Leaf emergence rate from branch apical bud (no. d–1) Branch stem elongation rate (mm d–1) Week 1 Week 3 Week 1 Week 3 P10 0.167 0.190 6.91 6.36 (0.0151) (0.0151) (0.275) (0.453) P14 0.071 0.076 1.12 1.21 (0.0184) (0.0216) (0.527) (0.750) Branch position Leaf emergence rate from branch apical bud (no. d–1) Branch stem elongation rate (mm d–1) Week 1 Week 3 Week 1 Week 3 P10 0.167 0.190 6.91 6.36 (0.0151) (0.0151) (0.275) (0.453) P14 0.071 0.076 1.12 1.21 (0.0184) (0.0216) (0.527) (0.750) At the start of the assessment period the root at P10 was excised and the number of leaves on, and lengths of, the P10 and P14 branches measured. These measurements were repeated after 1, 2, and 3 weeks. The mean daily rates of leaf emergence and stem elongation (±SE) were independently calculated (n=6). Open in new tab Table 1. Mean rates of leaf emergence on, and stem elongation of, the branches at P10 and P14 of treatment E during the first and third weeks of the assessment period of the Trifolium experiment Branch position Leaf emergence rate from branch apical bud (no. d–1) Branch stem elongation rate (mm d–1) Week 1 Week 3 Week 1 Week 3 P10 0.167 0.190 6.91 6.36 (0.0151) (0.0151) (0.275) (0.453) P14 0.071 0.076 1.12 1.21 (0.0184) (0.0216) (0.527) (0.750) Branch position Leaf emergence rate from branch apical bud (no. d–1) Branch stem elongation rate (mm d–1) Week 1 Week 3 Week 1 Week 3 P10 0.167 0.190 6.91 6.36 (0.0151) (0.0151) (0.275) (0.453) P14 0.071 0.076 1.12 1.21 (0.0184) (0.0216) (0.527) (0.750) At the start of the assessment period the root at P10 was excised and the number of leaves on, and lengths of, the P10 and P14 branches measured. These measurements were repeated after 1, 2, and 3 weeks. The mean daily rates of leaf emergence and stem elongation (±SE) were independently calculated (n=6). Open in new tab Effect of parent apical bud growth rates on the outgrowth of test buds in the core treatments (A–D): Comparison of responses of test buds formed on the faster growing branches at P10 with those formed on the slower growing P14 branches shows there to have been a clear delay in their start of outgrowth on the more distal branches (Fig. 3). This was so both at high NRS levels (nodal roots at P10 retained, A versus B) and at low levels (nodal roots at P10 excised, C versus D). For leaf emergence (Fig. 3a) this delay was about 7 d, but the delay in the start of bud stem elongation (Fig. 3b) was closer to 3 weeks. Once outgrowth had started, bud leaf emergence on branches at P14 was about 15% less than on P10 branches but, even 35 d after treatment, bud stem elongation on P14 branches was less than half that of equivalent buds on P10 branches. Effect of test bud positions on P10 and P14 branches on their outgrowth: Test buds on the branch at P10 in treatment C grew out faster than those at P14 in treatment D (Fig. 3). Treatment F [Fig.1a(iv) and 1b] was designed to test whether this difference was related to differences in ages of the branches at P10 and P14, and hence the position of test buds on them, rather than the activation levels (growth rates) of their apical buds. Comparison of the results of treatments F, C and D, in which test bud outgrowth after nodal root excision occurred at low NRS levels (Fig. 3), shows, however, that the observed differences between C and D were not the result of the position on branches of the test buds. In treatment F (with P14 nodally rooted during the pretreatment phase) test bud outgrowth was markedly higher than in treatment D that never possessed a root at P14, even though the position of test buds was the same in both (Fig. 1b). Comparison of treatments F and C, in each of which the branch bearing the test buds was borne at a previously rooted phytomer (for 26 d at P14 in F, compared with 45 d at P10 in C), showed that outgrowth of test buds in F was much closer to that of C than of D, indicating that the effect of the position of buds on branches was minor. (b) Hypothesis 2: outgrowth of an axillary bud depends upon both its outgrowth potential and the NRS supply to it Effect of net root-derived stimulus (NRS): Test buds either received a high supply of NRS when the nodal root system at P10 was retained (treatments A and B) or a lower supply, when the nodal root system at P10 was excised (treatments C and D) (Fig. 1b). Comparison of A with C and B with D (Fig. 3) shows that both leaf emergence and stem elongation of axillary buds were significantly higher, by 29% and 80%, respectively, by the end of the assessment period (P <0.001) when their NRS supply was higher. The ANOVA found that the interaction between NRS supply and branch position (outgrowth potential) was non-significant (P >0.05) for both leaf emergence and stem elongation. Thus NRS and outgrowth potential each independently influence test bud growth in an additive fashion. Tradescantia experiment: bud outgrowth responses on the primary stem This experiment tested Hypothesis 1 only: that outgrowth potential of an axillary bud is directly related to the growth rate (activation level) of its parent apical bud. Whole plant responses to rooting treatments: The dry weights of the dissected parts of the shoot systems determined at the conclusion of the experiment (Table 2) indicate that plants receiving the shortest (0D) and longest (24D) exposures to nodal roots (Fig. 2) did not differ in the weights of their large basal portions and that the difference between total shoot dry weights, including distal portions, was less than 10%. Table 2. Dry weights (g) of the basal portions of the shoot (primary stem and branches at phytomers 1–18) and the distal portions (phytomers 19–34) of Tradescantia fluminensis in treatments 0D and 24D at the end of the Tradescantia experiment; n=6 Exposure to nodal root (d) Basal Distal 0 25.86±3.578 0.97±0.297 24 26.21±2.990 3.47±0.449 Exposure to nodal root (d) Basal Distal 0 25.86±3.578 0.97±0.297 24 26.21±2.990 3.47±0.449 Open in new tab Table 2. Dry weights (g) of the basal portions of the shoot (primary stem and branches at phytomers 1–18) and the distal portions (phytomers 19–34) of Tradescantia fluminensis in treatments 0D and 24D at the end of the Tradescantia experiment; n=6 Exposure to nodal root (d) Basal Distal 0 25.86±3.578 0.97±0.297 24 26.21±2.990 3.47±0.449 Exposure to nodal root (d) Basal Distal 0 25.86±3.578 0.97±0.297 24 26.21±2.990 3.47±0.449 Open in new tab Effect of nodal roots on activation of primary stem apical buds: The duration of nodal root presence at node 19 had a significant (P <0.001) and continuing effect on primary stem elongation in the region immediately proximal to the apical buds (Fig. 4). Stem elongation was 50% greater in plants exposed to the influence of nodal roots for 24 d (treatment 24D) than in the non-rooted control plants (treatment 0D) by the end of Phase 1 (see Fig. 2 for details of Phases 1 and 2). Strong treatment effects were also observed on the rates of leaf emergence over the whole 9 week Phase 1 period and elongation of the primary stem during week nine (Table 3). Table 3. Mean rate of leaf emergence (no. d–1) on the primary stem during Phase 1, and mean elongation rate (mm d–1) of the primary stem during the last week of Phase 1 (week 9), for each treatment of the Tradescantia experiment, n=6, (±SE) Duration of exposure to nodal root (d) 0 8 12 18 24 Leaf emergence 0.162 0.196 0.227 0.235 0.241 (0.0426) (0.0278) (0.0259) (0.0186) (0.0119) Stem elongation 3.88 3.68 4.54 5.21 6.04 (1.167) (0.560) (0.490) (0.391) (0.449) Duration of exposure to nodal root (d) 0 8 12 18 24 Leaf emergence 0.162 0.196 0.227 0.235 0.241 (0.0426) (0.0278) (0.0259) (0.0186) (0.0119) Stem elongation 3.88 3.68 4.54 5.21 6.04 (1.167) (0.560) (0.490) (0.391) (0.449) Open in new tab Table 3. Mean rate of leaf emergence (no. d–1) on the primary stem during Phase 1, and mean elongation rate (mm d–1) of the primary stem during the last week of Phase 1 (week 9), for each treatment of the Tradescantia experiment, n=6, (±SE) Duration of exposure to nodal root (d) 0 8 12 18 24 Leaf emergence 0.162 0.196 0.227 0.235 0.241 (0.0426) (0.0278) (0.0259) (0.0186) (0.0119) Stem elongation 3.88 3.68 4.54 5.21 6.04 (1.167) (0.560) (0.490) (0.391) (0.449) Duration of exposure to nodal root (d) 0 8 12 18 24 Leaf emergence 0.162 0.196 0.227 0.235 0.241 (0.0426) (0.0278) (0.0259) (0.0186) (0.0119) Stem elongation 3.88 3.68 4.54 5.21 6.04 (1.167) (0.560) (0.490) (0.391) (0.449) Open in new tab Growth rates of test axillary buds during three weeks following decapitation of the primary stem: At the time of decapitation all test buds were still completely enclosed within the sheaths of their subtending leaves in all treatments and the decapitation treatment stimulated their outgrowth to varying degrees (Fig. 5). Duration of previous exposure to nodal roots during Phase 1 had significant (P <0.001) effects on the elongation of the test bud stems. Test buds on plants that had retained their nodal roots for longer during Phase 1 grew out sooner and formed faster growing branches than on those exposed to the influence of a nodal root for shorter periods (Fig. 5). In non-decapitated control plants that had been previously exposed to a root for 0, 8 or 12 d, there was no measurable outgrowth of the buds equivalent to X, Y, and Z by day 23 after treatment. In those that retained a nodal root for 18 d or 24 d during Phase 1, the three equivalent buds attained mean stem lengths of 1.7 (±1.56) and 6.9 (±3.11) mm, respectively, compared with 30.3 (±5.59) and 32.7 (±4.68) mm in the decapitated plants (Fig. 5). Correlation of growth rates of test axillary buds and primary stem apical buds There was a direct linear relationship (P <0.01) between the rates of elongation of primary stem apical buds during the week preceding decapitation and those of the test axillary buds during weeks two and three following decapitation (Fig. 6). Discussion Central to this study was the hypothesis (Hypothesis 1) that, at the time axillary buds emerge from their parent apical bud, they have an outgrowth potential that is determined by the activation level (growth rate) of that bud. A key requirement of the experimental design was therefore to be able to vary the activation levels of apical buds while as far as possible holding all other variables constant. To achieve this, nodal rooting treatments were imposed that led to different growth rates of the apical buds on the P10 and P14 branches (Table 1) in the Trifolium experiment and on the primary stems (Table 3; Fig. 4) in the Tradescantia experiment. Outgrowth responses of test axillary buds that were newly emerged from these apical buds were then measured on plants that had been manipulated so as to (i) provide uniform NRS supply to comparable test buds and (ii) remove possible variation in the inhibitory influence of apical buds by excising these immediately distal to the test buds. Uniformity of NRS supply for both experiments was attained by means of the shoot excision procedures detailed earlier. In Trifolium, these were designed to channel all the NRS available at P10 solely to the test axillary buds, either on the P10 or P14 branches (Fig. 1). Although these buds were of similar chronological age and phytomeric distance (number of nodes) from P10, the physical distances from the P10 node to the test buds varied appreciably, the stem lengths between the youngest test buds and the node at P10 being: 305 (±12.9) mm in treatments A and C, 202 (±5.0) mm in B and D, and 215 (±7.5) mm in F. However, the buds that responded most strongly were those on the branches at P10, in A and C, and these were actually further from their sources of NRS than those with the weakest responses on the branches at P14 in B and D. Thus, the effect of distance that NRS travels along stems was small relative to the effects of outgrowth potential on the growth of axillary buds, as can be seen by comparison of the responses of treatment F with B, C, and D (Fig. 3). In the Tradescantia experiment, in which no lateral branches were excised (Fig. 2), the branches at the base of the plants (P1–P18), that would have acted as the largest competing sinks for NRS, were similar for all treatments (Table 2). Variation induced in distal branch development by the differences in exposure to nodal roots at P19 amounted to <10%, and, moreover, it was the test buds in the treatment with longest exposure to nodal roots (24D), and greatest development of distal competitive sinks (Table 2), that grew most strongly. Thus variation in NRS supply to the test buds in Tradescantia was not a major factor driving the results obtained. The design of the Trifolium experiment was based on the assumption that differences in the positions of the test buds on the branches at P10 and P14 would have little influence on their outgrowth. Those on the P10 branch were situated at the sixth, seventh, and eighth nodes that emerged on the branch whereas those on the P14 branch were at the first three formed on it, at the first, second, and third nodes (Fig. 1). Because the larger P10 branch bore eight emerged leaves and the smaller P14 branch only three, and because those on the P14 branch, being the first-formed on the branch, were smaller (Wilman and Simpson, 1988; Hay et al., 1993) than leaves at the sixth, seventh, and eighth phytomers on the P10 branch, this assumption required testing. Treatment F was designed as a means of doing this. Comparison of the results of treatments F (rooted at P14) and C and D (rooted at P10) support the above-stated assumption: outgrowth of the buds on the P14 branch in treatment F was markedly greater than that of D and was approaching that of C (Fig. 3). This was so, despite there being differences in branch sizes and developmental maturity and in the time for which nodal roots were present. Nodal roots were present for 45 d at P10 in treatments C and D compared with only 26 d at P14 in treatment F. Clearly the difference between the outgrowth of buds in response to a common NRS supply in treatments C and D was almost entirely, and perhaps wholly, the result of differences in their intrinsic outgrowth potentials rather than differences in their positions on branches. The excision of the parent apical buds immediately distal to the test buds in both experiments ensured that any variations in outgrowth of test axillary buds, under the same conditions of NRS supply, could be ascribed solely to variation in intrinsic properties of the axillary buds themselves rather than to the continuing influence of their parent apical buds. Observed variations in outgrowth were thus clearly a result of a carry-over of the conditioning of axillary buds by the environment prevailing within their parent apical bud during their early development. This was particularly apparent in Tradescantia in which the youngest test buds (Z, Fig. 2) were not initiated within their parent apical buds in any treatment until after the excision of the nodal roots, at which time all buds became wholly dependent upon NRS supplied from comparable basal root systems. The results of both experiments strongly support the retention of Hypothesis 1: that each axillary bud exhibits an outgrowth potential that is directly related to the growth rate of its parent apical bud. In both, there was a close correlation between outgrowth rates of test buds and the activation levels of their parent apical buds that held true in relation to the measured growth of axillary buds on stems of different size, developmental maturity, and hierarchy within plants. In Trifolium the test buds on the P10 branch grew more rapidly in response to treatment than those on the P14 branch (Fig. 3), while in Tradescantia, there was a progressively increasing rate of growth of test buds as duration of exposure to the influence of nodal roots increased from treatment 0D through to 24D (Fig. 5). Regression analyses (Fig. 6) indicate that the rates of stem elongation of test axillary buds were correlated in a positive linear fashion with those of the parent apical buds in which they formed. A second hypothesis (Hypothesis 2), that the rate of outgrowth of an axillary bud depends upon the combination of its outgrowth potential and the supply of NRS available to it, was also strongly supported by the results of the Trifolium experiment. In this, supply of NRS during the assessment period was varied by either retaining the distal nodal root at P10 (high NRS) or excising it so that NRS was supplied solely from the basal roots (low NRS). Analysis of the results (Fig. 3) showed that both bud outgrowth potential and NRS supply significantly influenced leaf emergence and stem elongation of test axillary buds. These factors were additive in their effect, the interaction between the two being non-significant. These results raise the question as to how widespread the regulatory role of apical bud activation level might be among other species. Previous studies on a range of annual and short-lived perennial dicotyledonous species have led to the suggestion that the outgrowth potential of axillary buds might be related to the developmental stage and physiological activity of their parent apical bud (McDaniel and Hsu, 1976; Gould et al., 1987; Grbic and Bleecker, 2000). Because the changes in apical bud activity observed in these cases largely arose in parallel with the transition to flowering, it is difficult to separate influences associated with the development of reproductive maturity from those directly related to apical bud growth rates. The present findings, though, using plants maintained throughout in a strictly vegetative ‘steady developmental state’, have demonstrated that there is, indeed, a direct relationship between parent apical bud growth rates and the outgrowth potential of axillary buds newly emerged from them, at least in the two species studied. It remains unclear the degree to which such a relationship is widespread among angiosperms, but it does seem likely that it might be common among prostrate nodally-rooting perennial herbs. It is pertinent in this regard that, despite the evolutionarily wide separation of Trifolium repens and Tradescantia fluminensis, they behave similarly in response to nodal roots. In both, stimulatory signals derived from nodal roots dominate their control of branching. In both, also, full activation of axillary buds is only weakly reversible following exposure to the influence of nodal roots (Fig. 4; Thomas and Hay, 2007). Similar stimulatory responses to nodal roots have also been found to occur in all other prostrate nodally-rooting perennial herbs examined so far (Thomas and Hay, 2004). The additional understanding provided by the present investigation, in combination with knowledge of the pattern of intra-plant NRS distribution previously reported (Thomas and Hay, 2008a), will now serve as a basis for future work developing an improved model of the physiological regulation of branching by basal roots that might pertain to prostrate nodal-rooting herbs as a group. We thank Jocelyn Tilbrook and Rachael Sheridan for technical assistance and John Koolaard for statistical advice. 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This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.5/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details) © 2009 The Author(s).

Shetty, Nandini P.; Jensen, Jens D.; Knudsen, Anne; Finnie, Christine; Geshi, Naomi; Blennow, Andreas; Collinge, David B.; Jørgensen, Hans J. Lyngs

doi: 10.1093/jxb/erp269pmid: 19880540

Abstract The accumulation of the pathogenesis-related (PR) proteins β-1,3-glucanase and chitinase and structural defence responses were studied in leaves of wheat either resistant or susceptible to the hemibiotrophic pathogen Septoria tritici. Resistance was associated with an early accumulation of β-1,3-glucanase and chitinase transcripts followed by a subsequent reduction in level. Resistance was also associated with high activity of β-1,3-glucanase, especially in the apoplastic fluid, in accordance with the biotrophic/endophytic lifestyle of the pathogen in the apoplastic spaces, thus showing the highly localized accumulation of defence proteins in the vicinity of the pathogen. Isoform analysis of β-1,3-glucanase from the apoplastic fluid revealed that resistance was associated with the accumulation of an endo-β-1,3-glucanase, previously implicated in defence against pathogens, and a protein with identity to ADPG pyrophosphatase (92%) and germin-like proteins (93%), which may be involved in cell wall reinforcement. In accordance with this, glycoproteins like extensin were released into the apoplast and callose accumulated to a greater extent in cell walls, whereas lignin and polyphenolics were not found to correlate with defence. Treatment of a susceptible wheat cultivar with purified β-1,3-glucan fragments from cell walls of S. tritici gave complete protection against disease and this was accompanied by increased gene expression of β-1,3-glucanase and the deposition of callose. Collectively, these data indicate that resistance is dependent on a fast, initial recognition of the pathogen, probably due to β-1,3-glucan in the fungal cell walls, and this results in the accumulation of β-1,3-glucanase and structural defence responses, which may directly inhibit the pathogen and protect the host against fungal enzymes and toxins. ADPG pyrophosphatase, callose, chitinase, extensin, β-1,3-glucanase, Mycosphaerella graminicola, PAMPs/MAMPs, Septoria tritici, wheat Introduction The innate ability of plants to detect pathogens is essential for their survival (Nürnberger et al., 2004; Altenbach and Robatzek, 2007). This is made possible by the ability of the plant to recognize non-self structures through receptors (pattern recognition receptors, PRR) and the structures recognized by these receptors are termed pathogen-/microbe-associated molecular patterns or PAMPs/MAMPs (Nürnberger et al., 2004; Altenbach and Robatzek, 2007). Several different PAMPs have been isolated from bacteria, Oomycetes, and fungi. Examples from fungi include β-1,3-glucan, chitin, and ergosterol (Nürnberger et al., 2004; Altenbach and Robatzek, 2007). Pathogenesis-related (PR) proteins have been implicated in disease resistance in several plant–pathogen interactions (Linthorst, 1991; Van Loon and Van Strien, 1999). The roles of some PR-proteins are poorly understood (Kitajima and Sato, 1999; Van Loon and Van Strien, 1999), whereas others are known to have antifungal activity in vivo. Thus, the two plant hydrolases PR-2 (β-1,3-glucanase) and PR-3 (chitinase) degrade fungal cell walls and may therefore inhibit pathogen growth directly (Kim and Hwang, 1997; Kini et al., 2000). Oligomers of β-1,3-glucan and chitin released from the hydrolysis of fungal cell walls may, furthermore, act as elicitors of defence reactions (Takeuchi et al., 1990; Wu et al., 1997; Jia and Martin, 1999) and thus serve as PAMPs/MAMPs (Nürnberger et al., 2004; Altenbach and Robatzek, 2007). As a result of recognition of PAMPs/MAMPs, defence reactions are activated in the plant. These responses include the accumulation of Reactive Oxygen Species (ROS), and PR-proteins as well as the reinforcement of the cell wall by oxidative cross-linking of cell wall components and the deposition of callose and lignin (Nürnberger et al., 2004; Göhre and Robatzek, 2008). The synthesis of callose, a β-1,3-glucan, occurs de novo as a response to pathogen attack (Skou et al., 1984; Enkerli et al., 1997; Verma and Hong, 2001) although it is present at a constitutive level in many other regions such as pollen tubes (Meikle et al., 1991) and sieve tubes (Skou et al., 1984). Lignin precursors and other phenolics are themselves directly toxic to pathogens and their polymerization makes cell walls more difficult to penetrate and degrade (Hammerschmidt and Kuć, 1982). Likewise, cross-linking involving hydroxyproline-rich glycoproteins, including extensin may have the same effect (Wei and Shirsat, 2006). In addition to their role in the activation of defence response genes and direct antimicrobial effects, the accumulation of ROS is reported to enhance cross-linking in cell walls (Thordal-Christensen et al., 1997). Septoria tritici Roberge in Desmaz [teleomorph Mycosphaerella graminicola (Fuckel) J. Schröt. in Cohn] is a serious constraint for wheat (Triticum aestivum L.) production, causing Septoria tritici blotch or speckled leaf blotch. The disease has become more serious worldwide during recent years (Eyal, 1999). The infection biology of S. tritici is fairly well understood (Cohen and Eyal, 1993; Kema et al., 1996; Duncan and Howard, 2000; Shetty et al., 2003) whereas detailed studies of host defence responses against S. tritici and studies of how the pathogen influences host physiology have only recently begun (Ray et al., 2003; Shetty et al., 2003, 2007; Keon et al., 2007; Rudd et al., 2008). Cohen and Eyal (1993) observed callose to accumulate in a resistant, but not in a susceptible cultivar. However, they stated that there was no conclusive evidence for the involvement of callose in resistance. No evidence has been found that resistance was associated with compartmentalization (Kema et al., 1996), classical hypersensitive responses (HR) (Kema et al., 1996; Shetty et al., 2003; Rudd et al., 2008) or polyphenolic compounds or lignin (Cohen and Eyal, 1993; Kema et al., 1996), although Shetty et al. (2003) observed autofluorescence to occur to a significantly higher degree in a resistant than in a susceptible wheat cultivar. Recently, H2O2 accumulation was reported as a host response in wheat to infection by S. tritici and evidence was presented that H2O2 was indeed a factor inhibiting pathogen growth in wheat (Shetty et al., 2003, 2007). However, it was also concluded that H2O2 was probably not the only defence response since scavenging of H2O2 from a resistant cultivar did not render it fully susceptible to the pathogen (Shetty et al., 2007). It is shown here that PR-protein accumulation and callose deposition correlate with resistance in wheat against infection by S. tritici. Furthermore, it is shown that the application of purified β-1,3-glucan isolated from the cell walls of the pathogen is able to protect a susceptible wheat cultivar from disease development, and that this is accompanied by, among other defences, the accumulation of PR-proteins and callose. Thus our data suggest that β-1,3-glucanase cleaves β-1,3-glucan in the pathogen cell wall, releasing fragments which elicit further structural defence responses, to participate in preventing the colonization of S. tritici. Materials and methods Plants and inoculation Two wheat cultivars were used throughout the experiments: cv. Sevin is susceptible to isolate IPO323 of S. tritici and cv. Stakado is resistant to this isolate. Plants were grown, inoculum produced, and plants inoculated as described previously by Shetty et al. (2003). Control plants were treated with distilled water. RNA extraction and quantification of gene expression by quantitative real-time RT-PCR Leaves of S. tritici-inoculated or water-treated plants of both cultivars were sampled at 1, 3, 5, 7, 9, 11, 13, and 15 dai, ground in liquid nitrogen, and stored at –80 °C until use. Total RNA was extracted from 100 mg homogenized plant tissue using the RNeasy Plant Mini Kit (Qiagen, Venlo, The Netherlands) following the manufacturer's protocol. Genomic DNA contaminating the samples was removed by treatment with DNase using DNA-free (Ambion, UK) according to the manufacturer's instructions. cDNA synthesis was carried out using iscript (Bio-Rad, USA) according to the manufacturer's protocol. The final concentration of reverse transcribed total RNA was 35 ng μl−1. For each sample, a negative control was made without adding reverse transcriptase to ensure that there was no contamination with genomic DNA. The primers used are shown in Table 1. The 18S rRNA gene was used as the reference gene (Shimada et al., 2003). The other primers were designed using the primer3 program (http://fokker.wi.mit.edu/primer3/input.htm). Specificities of the genes were tested by blasting analyses of the amplicon. Tests for secondary structure of the amplicon was performed on the website: http://mfold.bioinfo.rpi.edu/. Testing of primers as well as real-time RT-PCR was carried out as described by Bedini et al. (2005). Table 1. Primers used in the quantitative real-time RT-PCR studies Gene Accession Forward primer Reverse primer 18S ribosomal RNA 5′-CGGCTACCACATCCAAGGAA-3′ 5′-GCTGGAATTACCGCGGCT-3′ β-1,3-glucanase DQ090946.1 5′-AACGACCAGCTCTCCAACAT-3′ 5′-GTATGGCCGGACATTGTTCT-3′ Chitinase AY437443.1 5′-ACGGTGTGATCACCAACATC-3′ 5′-CAGTCCAGGTTGTCACCGTA-3′ PAL AY005474.1 5′-CCAATGTTCTGTCCGTCCTT-3′ 5′-CTTCAGCTTGTGGGTCAGGT-3′ Chalcone synthase AY286097.1 5′-TCACCTTCCACCTCCTCAAG-3′ 5′-GGATGCGCTATCCAGAAGAC-3′ Oxalate oxidase AJ556991 5′-TGCAGTTCAACGTCGGTAAG-3′ 5′-ATGGCACGAAGACGATACC-3′ Gene Accession Forward primer Reverse primer 18S ribosomal RNA 5′-CGGCTACCACATCCAAGGAA-3′ 5′-GCTGGAATTACCGCGGCT-3′ β-1,3-glucanase DQ090946.1 5′-AACGACCAGCTCTCCAACAT-3′ 5′-GTATGGCCGGACATTGTTCT-3′ Chitinase AY437443.1 5′-ACGGTGTGATCACCAACATC-3′ 5′-CAGTCCAGGTTGTCACCGTA-3′ PAL AY005474.1 5′-CCAATGTTCTGTCCGTCCTT-3′ 5′-CTTCAGCTTGTGGGTCAGGT-3′ Chalcone synthase AY286097.1 5′-TCACCTTCCACCTCCTCAAG-3′ 5′-GGATGCGCTATCCAGAAGAC-3′ Oxalate oxidase AJ556991 5′-TGCAGTTCAACGTCGGTAAG-3′ 5′-ATGGCACGAAGACGATACC-3′ Open in new tab Table 1. Primers used in the quantitative real-time RT-PCR studies Gene Accession Forward primer Reverse primer 18S ribosomal RNA 5′-CGGCTACCACATCCAAGGAA-3′ 5′-GCTGGAATTACCGCGGCT-3′ β-1,3-glucanase DQ090946.1 5′-AACGACCAGCTCTCCAACAT-3′ 5′-GTATGGCCGGACATTGTTCT-3′ Chitinase AY437443.1 5′-ACGGTGTGATCACCAACATC-3′ 5′-CAGTCCAGGTTGTCACCGTA-3′ PAL AY005474.1 5′-CCAATGTTCTGTCCGTCCTT-3′ 5′-CTTCAGCTTGTGGGTCAGGT-3′ Chalcone synthase AY286097.1 5′-TCACCTTCCACCTCCTCAAG-3′ 5′-GGATGCGCTATCCAGAAGAC-3′ Oxalate oxidase AJ556991 5′-TGCAGTTCAACGTCGGTAAG-3′ 5′-ATGGCACGAAGACGATACC-3′ Gene Accession Forward primer Reverse primer 18S ribosomal RNA 5′-CGGCTACCACATCCAAGGAA-3′ 5′-GCTGGAATTACCGCGGCT-3′ β-1,3-glucanase DQ090946.1 5′-AACGACCAGCTCTCCAACAT-3′ 5′-GTATGGCCGGACATTGTTCT-3′ Chitinase AY437443.1 5′-ACGGTGTGATCACCAACATC-3′ 5′-CAGTCCAGGTTGTCACCGTA-3′ PAL AY005474.1 5′-CCAATGTTCTGTCCGTCCTT-3′ 5′-CTTCAGCTTGTGGGTCAGGT-3′ Chalcone synthase AY286097.1 5′-TCACCTTCCACCTCCTCAAG-3′ 5′-GGATGCGCTATCCAGAAGAC-3′ Oxalate oxidase AJ556991 5′-TGCAGTTCAACGTCGGTAAG-3′ 5′-ATGGCACGAAGACGATACC-3′ Open in new tab Extraction of apoplastic fluid Apoplastic fluid was obtained according to the method of Kerby and Somerville (1989). Leaves were sampled at 1, 3, 5, 7, 9, 11, 13, and 15 dai. At each time point, the leaves were immersed in deionized water and then vacuum infiltrated for about 60 min. After infiltration, they were gently dried with tissue paper, wrapped in a plastic film, and centrifuged at 500 g, at 4 °C with their cut end down in a centrifuge tube with glass beads at the bottom. The collected apoplastic fluids were immediately frozen at –80 °C until use. Assays for β-1,3-glucanase and chitinase activity Activity of the enzymes was assayed in apoplastic fluid and in protein extracts from whole leaves. For both types of extracts, protein was quantified in duplicate in an ELISA reader at 595 nm according to the method of Bradford (1976) using the Bio-Rad protein assay (Bio-Rad Ltd) with bovine albumin (Sigma) as standard. For whole leaf extracts, leaves were harvested at 1, 3, 5, 7, 9, 11, 13, and 15 dai, ground in liquid nitrogen and stored at –80 °C until use. Protein was extracted in 0.05 M sodium acetate buffer (pH 5.2) at 4 °C and the homogenate centrifuged at 10 000 g for 30 min. at 0 °C. The supernatant was used as the source of enzyme and stored on ice until analysis. β-1,3-glucanase activity was assayed according to Kini et al. (2000) with slight modifications. The samples were incubated with 0.1% (w/v) laminarin in 0.05 M sodium acetate buffer (pH 5.2). The mixture was incubated for 15 min. in a shaker at 37 °C. The reaction was stopped by adding DNS reagent [0.5% (w/v) 3,5-dinitrosalicylic acid (Sigma) and 15% (w/v) potassium sodium tartrate tetrahydrate (Sigma)] followed by boiling in a water bath for 10 min. The absorbance was measured in an ELISA-reader at 540 nm. A standard curve relating the amount of glucose equivalents to the absorbance at 540 nm was used to determine the activity. Chitinase activity was assayed according to Boller and Mauch (1988). The samples were incubated with colloidal chitin (made from chitin, Sigma) for 2 h at 37 °C. The reaction was stopped by centrifugation and the supernatant boiled with 0.8 M potassium borate buffer (pH 9.1) for 3 min. Subsequently, p-dimethylaminobenzaldehyde (Sigma) was added and the mixture incubated at 37 °C for 20 min, after which absorbance was measured at 585 nm in an ELISA-reader. A standard curve relating the amount of glucosamine equivalents to the absorbance at 585 nm was used to determine the activity. All enzyme activities are expressed as specific activity (μmol min−1 mg−1 protein). β-1,3-glucanase isoforms β-1,3-glucanase isoforms were determined by native-PAGE from apoplastic fluid according to the method of Schrauwen (1966). Apoplastic fluid was obtained as before from both Sevin and Stakado, either inoculated with S. tritici or treated with water at 1, 3, 5, 7, 9, and 11 dai (data not shown for water controls). Protein was quantified as before. Protein (40 μg) was electrophoresed in 10% polyacrylamide gels. Basic gels (pH 6.8) were used for the detection and separation of acidic isoforms and acidic gels (pH 5.2) for the separation of the basic isoforms. To detect the different isoforms of β-1,3-glucanase, the gel was stained for 30 min after electrophoresis in a solution containing 100 mg Laminarin in 25 ml 0.05 M Na-acetate buffer (pH 5.2) at 40 °C. After three washes, the gel was transferred to a glass tray containing 0.15% 2,3,5 triphenyl tetrazolium chloride (Sigma) in NaOH (150 mg in 20 ml 1 N NaOH) and boiled until red bands appeared. To check for equal loading of protein and for the identification of proteins by mass spectrometry, a gel was run as described above and stained with Coomassie Brilliant Blue. Mass spectrometry Gel pieces were excised from the Coomassie Brilliant Blue-stained bands. Protein was digested in-gel with sequencing grade porcine trypsin (Promega) as described by Zhang et al. (2007) and applied to a 600 μM AnchorChip™ target (Bruker Daltonics, Bremen, Germany) using a sample matrix wash procedure (Zhang et al., 2007) with α-hydroxycinammic acid as the matrix. MALDI-TOF (matrix assisted laser desorption ionization-time of flight) spectra were acquired using a Bruker Ultraflex II (Bruker Daltonics) in positive ion reflector mode. External calibration was performed using a tryptic digest of β-lactoglobulin. Spectra were processed in the FlexAnalysis software version 3.1 (Bruker Daltonics) and database searching was performed using BioTools version 3.0 (Matrix Science) to search the NCBI sequence database and the DFCI wheat gene index (http://compbio.dfci.harvard.edu/tgi/cgi-bin/tgi/gimain.pl?gudb=wheat) by applying the following criteria: taxonomy green plants; monoisotopic mass accuracy <80 ppm; one allowed missed cleavage site; carbamidomethylation of cysteine (complete), and oxidation of methionine (partial). The signal peptide cleavage sites were predicted using SignalP (Nielsen et al., 1997). Histochemical staining for callose, lignin, polyphenolic substances, and H2O2 Leaves were harvested 1, 3, 5, 7, and 9 days after inoculation (dai) and cleared using an ethanol:acetic acid mixture as previously described by Shetty et al. (2003). Callose deposition in cleared leaves was detected after staining the leaves with a solution of 0.005% Aniline Blue in 0.15 M dipotassium hydrogen phosphate (pH 8.2) for 2 h. The samples were observed using epifluorescence microscopy (excitation maximum 330–385 nm, dichroic mirror DM 400, barrier filter >420 nm). Regions with callose deposition emitted a greenish yellow fluorescence. For each time point, four leaves were examined and, on each leaf, 20 microscopic fields were studied (400× magnification, total area in field of vision approximately 0.22 mm2). The fields were selected randomly across the leaf. In each field, the total number of cells was counted as well as the number of cells with callose. These numbers also comprised cells of which only a part was seen within the field of vision. To study the distribution of callose inside the tissue, leaf pieces, 4×7 mm were cut from inoculated leaves and fixed in 2.5% glutaraldehyde in 0.1 M phosphate buffer (pH 6.8) under vacuum. After 24 h, the leaf pieces were washed in buffer and dehydrated in a graded series of isopropanol (IPA) and embedded in paraffin. After sectioning and drying of the slides, the paraffin was removed by UltraClear (Mallinckrodt Baker B.V. Devanter, Holland); and the sections transferred to 100% IPA. Finally, the slides were air-dried and mounted with Permount (Fisher Chemical, Fair Lawn, New Jersey, USA). Lignin accumulation was studied in cleared leaf pieces after staining with 2% phloroglucinol dissolved in 96% ethanol for 1 h. Subsequently, the tissue was placed on a glass slide with 10% HCl, heated slightly over a flame, and examined in the microscope. Pink to reddish staining of the xylem served as a control for the staining procedure. Polyphenolic substances were detected in cleared leaf pieces after staining with 0.05% Toluidine Blue O (pH 6.8) for 30 min and washing in phosphate buffer (pH 6.8). Turquoise staining of the xylem served as a control for the staining procedure. In vivo detection of H2O2 was carried out using the DAB-staining method as described before (Shetty et al., 2003). These leaves were also stained with Aniline Blue and examined for the presence of callose. Detection of cell wall polysaccharides in apoplastic fluid and leaves Leaves of cvs Sevin and Stakado were inoculated with S. tritici or treated with water as before and apoplastic fluid collected at 1, 3, 5, 7, 9, 11, 13, and 15 dai. Cell wall-derived polysaccharides in the apoplastic fluid and the leaves from which apoplastic fluid was extracted were analysed by immune-dot-assay and Western blot, respectively, using hybridoma supernatants of rat monoclonal antibodies LM1, LM2, LM5, LM6, LM7, JIM5, JIM7, and XGA2. These antibodies recognize specifically extensin, arabinogalactan, β-1,4-galactan, α-1,5-arabinan, homogalacturonan, sequentially methyl esterified homogalacturonan, non-sequentially methyl esterified homogalacturonan, and xylogalacturonan, respectively. These hybridoma supernatants were a kind gift from Professor P Knox (University of Leeds, UK). Immuno-dot-analysis followed the method described by Willats et al. (2002) using 1 μl of apoplastic fluid. Hybridoma supernatants and anti-rat secondary antibody conjugated with horseradish peroxidase were diluted in PBS containing 3% (w/v) skim milk in 1:10 and 1:1000, respectively. The remaining leaf materials, after isolating the apoplastic fluid (three leaves for each treatment; approximately 250 mg), were ground in 0.5 ml of 50 mM Na2CO3, 50 mM DTT, 15% (w/v) sucrose, and 2.5% (w/v) SDS using a glass homogenizer. Proteins were separated on a 12% SDS gel and analysed by Western blot using hybridoma supernatants and the secondary antibody diluted 1:40 and 1:1000, respectively. Preparation of β-glucan fragments from S. tritici mycelium To purify β-glucan from S. tritici, the fungus was grown in Fries medium for 3 weeks. After inoculation of the flasks, they were incubated in the dark for 48 h at 27 °C with continuous shaking (80 rpm). Subsequently, the flasks were incubated in darkness and without shaking at room temperature until use. After 3 weeks, the mycelial mat in the flasks was separated from the medium and freeze-dried. Water-soluble and mainly linear β-glucans were extracted from the freeze-dried mycelium (3 g dry matter) essentially as described by Yamaguchi et al. (2000). The dry mycelium was ground in a mortar with liquid N2. The fine (grey) powder was suspended in 50 ml PBS-buffer [0.1 M phosphate buffer (pH 7.2), 0.5 M NaCl] and the suspension was incubated for 15 min at room temperature. Following centrifugation, the pellet was recovered and re-extracted twice. The insoluble residues were further washed with 50 ml distilled water and then centrifuged followed by delipidation in four steps: (i) 50 ml acetone, (ii) 50 ml 1:1 ratio of chloroform:methanol (v/v), (iii) 50 ml chloroform, and (iv) 50 ml acetone. For each treatment, the insoluble matter was mixed thoroughly, incubated for 15 min at room temperature, and the insoluble residue was recovered by centrifugation. The final pellet was resuspended in 50 ml milliQ water and autoclaved three times for 15 min at 121 °C. The remaining insoluble residues were extracted overnight at room temperature with 50 ml 1 M NaOH containing 0.01% NaBH4. The pellet was re-extracted twice with 1 M NaOH and once with milliQ water. All supernatants were pooled and the mainly linear β-1,3-glucan (Yamaguchi et al., 2000) was precipitated by neutralization to pH 6.0 with glacial acetic acid and collected by centrifugation at 4 °C according to the protocol of Yamaguchi et al. (2000). Preparation of β-glucan fragments was achieved by partial enzymatic digestion using (endo-1,3-β-D-glucanase (Trichoderma sp.) EC 3.2.1.39). A total of 100 mg pure β-glucan was dispersed in 3 ml of 20 mM TRIS-HCl buffer (pH 4.5) and digested for 4.5 h at 40 °C with 0.5 U endo-β-1,3-glucanase (Megazyme). The reaction was stopped by boiling and the products recovered freeze-dried. The final product was analysed using High Performance Anion Exchange Chromatography (HPAEC) with pulsed amperometric detection (PAD) using a Dionex BioLC-System (Dionex Corp., Sunnyvale, CA) equipped with a CarboPac PA-200 column operating with a 0.5 ml ml−1 flow and an elution profile as described by Blennow et al. (1998). Chromatograms were visualized using the Chromeleon V.6.40 software. Application of purified β-glucan to plants In order to study the influence of purified β-glucan on the host, a range of experiments were carried out in cv. Sevin. Symptom expression of S. tritici was scored in plants either infiltrated (Shetty et al., 2007) or sprayed until run-off with a solution containing 100 μg purified β-glucan ml−1 of milliQ water. Plants were inoculated with S. tritici 24 h later as before and symptoms recorded 15 dai. Callose accumulation was studied as before in plants either sprayed with 100 μg purified β-glucan ml−1 of milliQ water or water (control), followed by inoculation with the pathogen. Observations were made at 3 dai and 5 dai. In order to test whether β-glucan application to the plants elicited gene expression, qRT-PCR were performed on plants either sprayed with purified β-glucan (100 μg ml−1) or water followed by inoculation with the pathogen. Samples were taken at 3 dai as before. The primers tested were PAL, chitinase, chalcone synthase, β-1,3-glucanase, and oxalate oxidase (Table 1). To examine the ability of the wheat plant to produce β-1,3-glucanase, which could degrade fungal cell walls, apoplastic fluid was isolated from cvs Stakado and Sevin (either inoculated with S. tritici or treated with water) at 3, 5, and 7 dai. The apoplastic fluid was used as the source of enzyme and purified fungal cell walls as substrate. Protein determination and β-1,3-glucanase assay was performed as described above. Statistical analysis Data from enzyme activity assays represent continuous variables and were analysed by analysis of variance assuming a normal distribution. Variances were stabilized by appropriate transformation of data if necessary. Data from studies of callose accumulation represent a discrete variable and were hence analysed by logistic regression, assuming a binomial distribution, as described by Shetty et al. (2003). All data were analysed by PC-SAS (release 8.2, SAS Institute, Cary, NC). For gene expression studies, statistical evaluations of the relative expression level of the target genes was evaluated in S. tritici-inoculated material compared with water-treated material and normalized to the 18S rRNA expression level. The analyses were performed using the relative expression software tool REST© as described by Pfaffl et al. (2002). The stability of the housekeeping gene was evaluated using the program Bestkeeper as described by Pfaffl et al. (2004). All experiments were repeated at least twice with similar results and representative results are presented. All hypotheses were rejected at P ≤0.05. In the following, all differences are significant at P ≤0.05 unless specifically mentioned. Results Gene expression Table 2 shows the results on gene expression. β-1,3-glucanase transcript levels in Stakado were elevated in inoculated compared with water-treated control plants at 1–3 dai and again at 9–11 dai, with no significant difference at 5–7 dai and a reduction at 13–15 dai. In Sevin, there was no significant difference between inoculated and control plants at 1–7 dai whereas elevated transcript levels were seen in inoculated plants from 9–15 dai. Chitinase transcript levels were elevated in Stakado in inoculated compared with control plants at 3–5 and 9 dai, whereas no significant differences were seen at 1, 7, and 13 dai. Reduced transcript levels were seen at 11 and 15 dai. In Sevin, no significant differences in transcript levels were observed at 1 and 5–7 dai, whereas reduced transcript levels were observed at 3 dai and elevated levels at 9–15 dai. PAL transcript levels were elevated at 3 and again at 9–11 and 15 dai in Stakado, whereas there were no significant differences between inoculated and control plants at the remaining time points. In Sevin, there was no significant difference between inoculated and control plants at 1–7 dai, whereas reduced transcript levels were seen at 9–11 dai followed by elevated levels at 13–15 dai. Table 2. Quantitative real-time RT-PCR experiments of transcript levels of β-1,3-glucanase, chitinase, PAL, and chalcone synthase gene expression in wheat cvs Stakado (resistant) and Sevin (susceptible) after inoculation with S. tritici Time β-1,3-Glucanase (PR-2)a Chitinase (PR-3)a PALa Chalcone synthasea Stakado Sevin Stakado Sevin Stakado Sevin Stakado Sevin 1 dai 4.5 A 1.4 NS 2.6 NS 0.9 NS 1.3 NS 0.8 NS 2.0 NS 0.6 NS 3 dai 4.1 A 1.1 NS 3.2 A 0.6 A 2.5 A 1.7 NS 0.4 A 1.2 NS 5 dai 2.9 NS 1.5 NS 1.6 A 1.5 NS 1.4 NS 0.8 NS 4.2 NS 0.6 NS 7 dai 1.0 NS 1.1 NS 1.0 NS 0.9 NS 1.1 NS 0.7 NS 0.7 A 0.5 NS 9 dai 3.5 A 52.0 A 5.3 A 54.9 A 2.3 A 0.5 A 0.6 A 0.1 A 11 dai 2.9 A 48.8 A 0.5 A 3.3 A 3.7 A 0.2 A 0.1 NS 0.1 A 13 dai 0.6 A 27.2 A 0.7 NS 317.4 A 1.1 NS 4.5 A 0.6 A 0.3 A 15 dai 0.5 A 322.9 A 0.3 A 2846.6 A 1.8 A 3.4 A 0.4 A 0.0 A,B Time β-1,3-Glucanase (PR-2)a Chitinase (PR-3)a PALa Chalcone synthasea Stakado Sevin Stakado Sevin Stakado Sevin Stakado Sevin 1 dai 4.5 A 1.4 NS 2.6 NS 0.9 NS 1.3 NS 0.8 NS 2.0 NS 0.6 NS 3 dai 4.1 A 1.1 NS 3.2 A 0.6 A 2.5 A 1.7 NS 0.4 A 1.2 NS 5 dai 2.9 NS 1.5 NS 1.6 A 1.5 NS 1.4 NS 0.8 NS 4.2 NS 0.6 NS 7 dai 1.0 NS 1.1 NS 1.0 NS 0.9 NS 1.1 NS 0.7 NS 0.7 A 0.5 NS 9 dai 3.5 A 52.0 A 5.3 A 54.9 A 2.3 A 0.5 A 0.6 A 0.1 A 11 dai 2.9 A 48.8 A 0.5 A 3.3 A 3.7 A 0.2 A 0.1 NS 0.1 A 13 dai 0.6 A 27.2 A 0.7 NS 317.4 A 1.1 NS 4.5 A 0.6 A 0.3 A 15 dai 0.5 A 322.9 A 0.3 A 2846.6 A 1.8 A 3.4 A 0.4 A 0.0 A,B Values shown represent fold up-regulation in inoculated compared with water-treated plants, after normalization to 18S rRNA (Shimada et al., 2003). a A, significant change; NS, non-significant change; B, exact value is 0.017. Open in new tab Table 2. Quantitative real-time RT-PCR experiments of transcript levels of β-1,3-glucanase, chitinase, PAL, and chalcone synthase gene expression in wheat cvs Stakado (resistant) and Sevin (susceptible) after inoculation with S. tritici Time β-1,3-Glucanase (PR-2)a Chitinase (PR-3)a PALa Chalcone synthasea Stakado Sevin Stakado Sevin Stakado Sevin Stakado Sevin 1 dai 4.5 A 1.4 NS 2.6 NS 0.9 NS 1.3 NS 0.8 NS 2.0 NS 0.6 NS 3 dai 4.1 A 1.1 NS 3.2 A 0.6 A 2.5 A 1.7 NS 0.4 A 1.2 NS 5 dai 2.9 NS 1.5 NS 1.6 A 1.5 NS 1.4 NS 0.8 NS 4.2 NS 0.6 NS 7 dai 1.0 NS 1.1 NS 1.0 NS 0.9 NS 1.1 NS 0.7 NS 0.7 A 0.5 NS 9 dai 3.5 A 52.0 A 5.3 A 54.9 A 2.3 A 0.5 A 0.6 A 0.1 A 11 dai 2.9 A 48.8 A 0.5 A 3.3 A 3.7 A 0.2 A 0.1 NS 0.1 A 13 dai 0.6 A 27.2 A 0.7 NS 317.4 A 1.1 NS 4.5 A 0.6 A 0.3 A 15 dai 0.5 A 322.9 A 0.3 A 2846.6 A 1.8 A 3.4 A 0.4 A 0.0 A,B Time β-1,3-Glucanase (PR-2)a Chitinase (PR-3)a PALa Chalcone synthasea Stakado Sevin Stakado Sevin Stakado Sevin Stakado Sevin 1 dai 4.5 A 1.4 NS 2.6 NS 0.9 NS 1.3 NS 0.8 NS 2.0 NS 0.6 NS 3 dai 4.1 A 1.1 NS 3.2 A 0.6 A 2.5 A 1.7 NS 0.4 A 1.2 NS 5 dai 2.9 NS 1.5 NS 1.6 A 1.5 NS 1.4 NS 0.8 NS 4.2 NS 0.6 NS 7 dai 1.0 NS 1.1 NS 1.0 NS 0.9 NS 1.1 NS 0.7 NS 0.7 A 0.5 NS 9 dai 3.5 A 52.0 A 5.3 A 54.9 A 2.3 A 0.5 A 0.6 A 0.1 A 11 dai 2.9 A 48.8 A 0.5 A 3.3 A 3.7 A 0.2 A 0.1 NS 0.1 A 13 dai 0.6 A 27.2 A 0.7 NS 317.4 A 1.1 NS 4.5 A 0.6 A 0.3 A 15 dai 0.5 A 322.9 A 0.3 A 2846.6 A 1.8 A 3.4 A 0.4 A 0.0 A,B Values shown represent fold up-regulation in inoculated compared with water-treated plants, after normalization to 18S rRNA (Shimada et al., 2003). a A, significant change; NS, non-significant change; B, exact value is 0.017. Open in new tab Chalcone synthase levels in Stakado were generally reduced (3, 7–9, and 13–15 dai) whereas no significant differences in levels were observed at the remaining time points. In Sevin (Table 2), there were no significant differences between inoculated and control plants 1–7 dai, followed by reduced transcript levels in inoculated plants 9–15 dai. Activity of β-1,3-glucanase and chitinase in whole leaf extracts and apoplastic fluid β-1,3-glucanase activity in whole leaf extracts (Fig. 1A) was higher in Stakado than in Sevin at all time points, with an increasing trend, especially from 11 dai. Activity in Stakado was higher in inoculated than in control plants at 5 dai and 13–15 dai, whereas the opposite relationship was observed at 3 dai. In apoplastic fluid (Fig. 1B), activity was also higher in Stakado than in Sevin, with very high activities observed in Stakado at 1 dai with a minor peak at 9 dai. At all time points, activities were higher in inoculated than in control plants. In Sevin, activity was higher in inoculated plants compared with the controls at 5–9 dai. Fig. 1. Open in new tabDownload slide Time-course of activity of β-1,3-glucanase (A, B) and chitinase (C, D) in Stakado and Sevin with and without inoculation with Septoria tritici, isolate IPO 323. (A) and (C) show activity in whole leaf extracts for β-1,3-glucanase and chitinase, respectively, whereas (B) and (D) show activity in the apoplastic fluid. Each value is presented ±standard error. Fig. 1. Open in new tabDownload slide Time-course of activity of β-1,3-glucanase (A, B) and chitinase (C, D) in Stakado and Sevin with and without inoculation with Septoria tritici, isolate IPO 323. (A) and (C) show activity in whole leaf extracts for β-1,3-glucanase and chitinase, respectively, whereas (B) and (D) show activity in the apoplastic fluid. Each value is presented ±standard error. Chitinase activity in whole leaf extracts (Fig. 1C) fluctuated considerably, often with no major differences between the inoculated and control plants. However, activity was higher in Stakado than in Sevin at 9 dai and 13–15 dai whereas higher activity was observed in Sevin at 1–3 dai. In apoplastic fluid (Fig. 1D), activity was considerably lower than in whole leaf extracts. Activity was higher in Stakado than in Sevin until 13 dai where activity in Sevin (inoculated with S. tritici) increased substantially. In Stakado, activity was higher in inoculated than in control plants at all time points, whereas in Sevin, no difference was seen between treatments until 13 dai after which time activity was higher in the inoculated plants. β-1,3-glucanase isoforms and identification by mass spectrometry Native-PAGE of apoplastic fluid showed the presence of several bands containing β-1,3-glucanase activity (Fig. 2B). The most intensely β-1,3-glucanase-stained band (Fig. 2, band b) was observed at all time points whereas another band (Fig. 2, band a) accumulated differentially, with high intensity at 3 dai and 7–11 dai. These bands were excised from gels and subjected to tryptic digestion and MALDI-TOF-TOF mass spectrometry. Identifications were performed using samples from Coomassie blue-stained gels, but similar spectra were obtained from the bands excised from activity-stained gels. For band b, database searches revealed a significant match (P ≤0.05) to a β-1,3-glucanase sequence from wheat (GenBank accession AAY96422.1), based on nine peptide masses with mass errors <20 ppm and covering 39% of the protein sequence. An additional peptide with [M+H]=2418.2 matched the N-terminus of the sequence after removal of the predicted signal peptide. Fig. 2. Open in new tabDownload slide Native-PAGE of β-1,3-glucanase isoforms accumulating in the apoplastic fluid of cv. Stakado (incompatible interaction) at 1, 3, 5, 7, 9, and 11 d after inoculation with S. tritici. (A) Coomassie Brilliant Blue stained gel and (B) activity stained gel. Band a was identified by mass spectrometry as adenosine diphosphate glucose pyrophosphatase and band b as endo-β-1,3-glucanase. Fig. 2. Open in new tabDownload slide Native-PAGE of β-1,3-glucanase isoforms accumulating in the apoplastic fluid of cv. Stakado (incompatible interaction) at 1, 3, 5, 7, 9, and 11 d after inoculation with S. tritici. (A) Coomassie Brilliant Blue stained gel and (B) activity stained gel. Band a was identified by mass spectrometry as adenosine diphosphate glucose pyrophosphatase and band b as endo-β-1,3-glucanase. The MS data for band a matched a wheat consensus sequence TC232338 which was highly similar to adenosine diphosphate glucose pyrophosphatase from Hordeum vulgare ssp. vulgare (93% identity to GenBank Accession CAC32847.1) and to germin-like protein 2a (92% identity to GenBank Accession ABG46233.1 also from Hordeum vulgare ssp. vulgare. Identification was based on three peptides with mass errors <50 ppm and covering 36% of the protein sequence. The identity of all peptides was confirmed by MSMS. The signal peptide cleavage site was confirmed by identification of the peptide LTQDFCVADLACPDTPAGYPCKK. Localization of callose, lignin, and polyphenolics The deposition of callose as a response to inoculation by S. tritici was observed in both resistant and susceptible wheat cultivars but at different rates and amounts (Table 3). Thus, deposition of callose in Stakado started 3 dai and increased in amount and intensity. Initially, callose was observed in and around stomata, (Fig. 3A, B), but at 5 dai, callose started to appear in the mesophyll beneath the stomata (Fig. 3C, D), coinciding with pathogen penetration and its confinement here. At 9 dai, callose was also seen in places far from substomatal cavities (Fig. 3E, F). In Sevin, deposition of callose started only 7 dai (Table 3). Here, deposition was also primarily seen in the stomatal complexes, with only a few reactions in the mesophyll. However, deposition was seen in very few cells and the intensity was very faint when compared to Stakado (data not shown). Table 3. Percentage of leaf cells with accumulation of callose in cvs. Stakado and Sevin after inoculation with Septoria tritici Stakado Sevin Odds ratioa 1 dai 0.0 0.0 1.00 NS 3 dai 0.4 0.0 ∞ * 5 dai 4.4 0.0 ∞ *** 7 dai 5.9 0.2 26.07 *** 9 dai 11.2 0.3 46.49 *** Stakado Sevin Odds ratioa 1 dai 0.0 0.0 1.00 NS 3 dai 0.4 0.0 ∞ * 5 dai 4.4 0.0 ∞ *** 7 dai 5.9 0.2 26.07 *** 9 dai 11.2 0.3 46.49 *** a Odds ratio for comparison of Stakado and Sevin (Sevin used as a reference, odds ratio=1.00). The number of asterisks indicates the degree of significance. NS, non-significant difference; *** significant at P≤0.001; * significant at P≤0.05. Open in new tab Table 3. Percentage of leaf cells with accumulation of callose in cvs. Stakado and Sevin after inoculation with Septoria tritici Stakado Sevin Odds ratioa 1 dai 0.0 0.0 1.00 NS 3 dai 0.4 0.0 ∞ * 5 dai 4.4 0.0 ∞ *** 7 dai 5.9 0.2 26.07 *** 9 dai 11.2 0.3 46.49 *** Stakado Sevin Odds ratioa 1 dai 0.0 0.0 1.00 NS 3 dai 0.4 0.0 ∞ * 5 dai 4.4 0.0 ∞ *** 7 dai 5.9 0.2 26.07 *** 9 dai 11.2 0.3 46.49 *** a Odds ratio for comparison of Stakado and Sevin (Sevin used as a reference, odds ratio=1.00). The number of asterisks indicates the degree of significance. NS, non-significant difference; *** significant at P≤0.001; * significant at P≤0.05. Open in new tab Fig. 3. Open in new tabDownload slide Deposition of callose in Stakado after inoculation with S. tritici isolate IPO323 (incompatible interaction). (A) Callose deposition in stoma in whole leaf mount 5 dai. (B) Deposition of callose in the guard cells in a transverse leaf section 5 dai. (C) Callose deposition in the mesophyll near a penetrated stoma 7 dai. (D) Callose deposition in the mesophyll near a penetrated stoma in a transverse section of a leaf 7 dai. (E, F) Callose deposition at sites remote from penetrated stomata in whole leaf mount (E) and in transverse leaf section (F) 9 dai. (G, H) Callose accumulation in leaves after pretreatment with water (G) or β-1,3-glucan (H) followed by inoculation with S. tritici at 5 dai (H). Arrows show deposition of callose. (This figure is available in colour at JXB online.) Fig. 3. Open in new tabDownload slide Deposition of callose in Stakado after inoculation with S. tritici isolate IPO323 (incompatible interaction). (A) Callose deposition in stoma in whole leaf mount 5 dai. (B) Deposition of callose in the guard cells in a transverse leaf section 5 dai. (C) Callose deposition in the mesophyll near a penetrated stoma 7 dai. (D) Callose deposition in the mesophyll near a penetrated stoma in a transverse section of a leaf 7 dai. (E, F) Callose deposition at sites remote from penetrated stomata in whole leaf mount (E) and in transverse leaf section (F) 9 dai. (G, H) Callose accumulation in leaves after pretreatment with water (G) or β-1,3-glucan (H) followed by inoculation with S. tritici at 5 dai (H). Arrows show deposition of callose. (This figure is available in colour at JXB online.) Staining of cleared leaf segments with phloroglucinol for the localization of lignin or Toluidine Blue O for the localization of polyphenolic substances did not reveal any accumulation related to pathogen growth in either cultivar at either time point. For both stains, positive reactions in the xylem showed that the staining procedure worked satisfactorily (data not shown). Detection of various cell wall polysaccharides in apoplastic fluid and leaves All antibodies tested showed similar patterns in apoplastic fluid. Figure 4 shows an example of LM1, which recognizes glycan components of a hydroxyproline-rich glycoprotein, extensin. In cv. Sevin, the LM1 epitope was relatively low in the apoplastic fluid regardless of the treatment, whereas it was associated well to the cell wall. In Stakado, the LM1 epitope was rather high during the early stages of the interaction (1–9 dai) in the inoculated samples, and particularly high at 11 dai. In the remaining material after collecting apoplastic fluid, the LM1 epitope in Sevin increased by 7–9 dai, whereas it decreased in Stakado in the inoculated samples, thus indicating that it is dissociated from the cell wall and flows out to the apoplastic fluid after inoculation. Fig. 4. Open in new tabDownload slide Detection of cell wall-derived glycoproteins in apoplastic fluid and in the leaves from which apoplastic fluid was isolated. (A) Apoplastic fluid was spotted onto a nitrocellulose membrane and extensin was detected by the LM1 antibody. (B) The remaining leaf material after isolation of apoplastic fluid at 7 and 9 dai was extracted and extensin was analysed by Western blotting using the LM1 antibody. Abbreviations: Sv323, Sevin inoculated with S. tritici; SvH2O, Sevin treated with water; St323, Stakado inoculated with S. tritici; StH2O, Stakado treated with water. (This figure is available in colour at JXB online.) Fig. 4. Open in new tabDownload slide Detection of cell wall-derived glycoproteins in apoplastic fluid and in the leaves from which apoplastic fluid was isolated. (A) Apoplastic fluid was spotted onto a nitrocellulose membrane and extensin was detected by the LM1 antibody. (B) The remaining leaf material after isolation of apoplastic fluid at 7 and 9 dai was extracted and extensin was analysed by Western blotting using the LM1 antibody. Abbreviations: Sv323, Sevin inoculated with S. tritici; SvH2O, Sevin treated with water; St323, Stakado inoculated with S. tritici; StH2O, Stakado treated with water. (This figure is available in colour at JXB online.) Application of purified β-glucan to plants In order to test more specifically whether β-glucans, which are potentially released from S. tritici as an effect of induced β-1,3-glucanase activity in wheat, mainly linear β-glucans were prepared from mycelium of the pathogen and fragmented by endo-1,3-β-D-glucanase from Trichoderma sp. The chain structure of the generated β-glucan fragments was analysed by High Performance Anion Exchange Chromatography (HPAEC) with pulsed amperometric detection (PAD), using linear malto oligosaccharides (linear α-1,4 glucans) and cellobiose (β-1,4) as standards (Fig. 5). The position of the peaks indicates that the preparation was composed of linear β-1,3-cello oligosaccharides with an apparent degree of polymerization (DP) ranging from 2 to 7. The main product is attributed to β-1,3-linked cellotriose, followed by longer cello oligosaccharides. Only very minor peaks of long fragments eluting at 15–16 min (approximately DP12) were detected (due to scaling, these are not visible in Fig. 5). Both spraying and infiltration of the purified β-glucan on the host resulted in complete inhibition of symptom expression at 15 dai whereas symptoms appeared in control plants treated with water. Figure 6 shows plants sprayed with either water or β-glucan. Fig. 5. Open in new tabDownload slide HPAEC-PAD profile for the linear β-1,3-cello oligosaccharides. Elution time for standard glucose (glc) and larger malto oligosaccharides (linear α-1,4-glucans, M2–M7) and cellobiose (β-1,4, C2) are indicated to show approximate fragment sizes. The large peak at 2 min is eluted salt. Fig. 5. Open in new tabDownload slide HPAEC-PAD profile for the linear β-1,3-cello oligosaccharides. Elution time for standard glucose (glc) and larger malto oligosaccharides (linear α-1,4-glucans, M2–M7) and cellobiose (β-1,4, C2) are indicated to show approximate fragment sizes. The large peak at 2 min is eluted salt. Fig. 6. Open in new tabDownload slide Spraying of wheat cv. Sevin with (A) water or (B) purified β-glucan from S. tritici followed by inoculation with the pathogen 24 h later. Symptoms scored 15 dai. (This figure is available in colour at JXB online.) Fig. 6. Open in new tabDownload slide Spraying of wheat cv. Sevin with (A) water or (B) purified β-glucan from S. tritici followed by inoculation with the pathogen 24 h later. Symptoms scored 15 dai. (This figure is available in colour at JXB online.) Application of a desalted protease and amylase-treated β-glucan preparation to remove possible protein or glycogen contamination (data not shown) showed that less β-glucan elicitor was required to obtain the identical effects on symptom expression. This indicates the presence of some impurities in the β-glucan preparation, but that these impurities had no or only a minor effect on the ability to protect against disease. Callose accumulation was studied as before at 3 dai and 5 dai in plants sprayed with β-glucan and it was found here that there was an increased accumulation of callose after β-glucan treatment (Fig. 3G, H). β-glucan application also affected gene expression. Thus, at 3 dai, chalcone synthase transcript levels were reduced (P <0.001), in β-1,3-glucan-treated compared to water-treated control plants (0.2-fold), whereas chitinase and β-1,3-glucanase levels were elevated (2.4- and 5.9-fold, respectively). There was no significant alteration of the expression of oxalate oxidase and PAL genes (1.2- and 1.3-fold, respectively). The ability of the wheat plant to produce β-1,3-glucanase, which could degrade fungal cell walls, was studied from apoplastic fluid isolated from cvs Stakado (Fig. 7) and Sevin (data not shown) either inoculated with S. tritici or treated with water. Figure 7 shows that apoplastic fluid isolated from Stakado at 5 d and 7 d after inoculation with S. tritici was better in degrading β-glucan from the cell walls of S. tritici than fluid isolated from plants sprayed with water. On the other hand, fluid from control plants was better in degrading fungal β-glucans than fluid from inoculated plants at 3 dai. Fig. 7. Open in new tabDownload slide Time-course study of activity of β-1,3-glucanase using apoplastic fluid from cv. Stakado as the source of enzyme and purified fungal cell walls from isolate IPO323 as the substrate. Apoplastic fluid was isolated from Stakado either inoculated with S. tritici or sprayed with water (controls) at different time points after inoculation (3, 5, and 7 dai). Each value is presented ±standard error. Comparisons are possible between inoculated and control plants at each time point. ** Significant at P ≤0.01, * Significant at P ≤0.05. Fig. 7. Open in new tabDownload slide Time-course study of activity of β-1,3-glucanase using apoplastic fluid from cv. Stakado as the source of enzyme and purified fungal cell walls from isolate IPO323 as the substrate. Apoplastic fluid was isolated from Stakado either inoculated with S. tritici or sprayed with water (controls) at different time points after inoculation (3, 5, and 7 dai). Each value is presented ±standard error. Comparisons are possible between inoculated and control plants at each time point. ** Significant at P ≤0.01, * Significant at P ≤0.05. Discussion After sensing an invading pathogen, plants activate a wide variety of general defence reactions including the oxidative burst, structural cell wall modifications, and the production of defence-related compounds such as PR-proteins (e.g. chitinases and β-1,3-glucanases) (Bolwell, 1999; Kini et al., 2000; Shetty et al., 2008). These proteins can degrade the cell walls of pathogens and inhibit their growth (Kim and Hwang, 1997). Furthermore, these enzymes hydrolyse fungal call walls, releasing β-1,3-glucan and chitin oligomers that act as elicitors of defence reactions (Takeuchi et al., 1990; Wu et al., 1997; Jia and Martin, 1999). Our data indicate that the PR-protein β-1,3-glucanase (PR-2) in wheat is essential for cleaving the cell walls of the hemibiotrophic pathogen Septoria tritici to release elicitors which can act as PAMPs to elicit further defence responses, which prevent colonization of the pathogen. Therefore, a series of experiments was conducted to elucidate the role of β-1,3-glucanase and β-1,3-glucan in defence. In the resistant cv. Stakado, elevated transcript levels were seen early in the interaction compared with the water-treated control, with a reduction during the late stages. In contrast, in the susceptible cv. Sevin, elevated gene expression was seen only from 9 dai. This change occurred before visible leaf necrosis occurred and probably coincides with the pathogen entering its necrotrophic phase (Shetty et al., 2003, 2007). Activity of β-1,3-glucanase was higher in Stakado than in Sevin, especially in apoplastic fluid, where very high levels were seen at 1 dai with the pathogen, coinciding with initial penetration and colonization of the host, which takes place in the apoplastic space (Shetty et al., 2003). This suggest that β-1,3-glucanase plays a role in the restriction of the pathogen. Since the pathogen lives as a biotroph/endophyte in the apoplastic spaces, this shows the highly localized accumulation of the defence protein close to the pathogen. On the other hand, activity of chitinase did not correlate well with resistance in the host or the infection course of the pathogen. Furthermore, the activity was very low in the apoplastic fluid. To investigate the potential involvement of β-1,3-glucanase in defence further, the temporal accumulation of different isoforms was studied. Native-PAGE analysis followed by β-1,3-glucanase staining revealed that, in apoplastic fluid from cv. Stakado, two intensely stained bands were present. By mass spectrometry, the band with the highest activity was identified as β-1,3-glucanase and this accession was identical to the accession tested in the transcript accumulation studies (Tables 1, 2). This β-1,3-glucanase has been implicated in defence against Puccinia striiformis in wheat (http://www.ncbi.nlm.nih.gov/nuccore/DQ090946.1) and also showed a prominent accumulation in the resistant cv. Stakado (Fig. 2), in compliance with the high β-1,3-glucanase activity (Fig. 1) and the high level of resistance to fungal colonization. The other band showed weaker β-1,3-glucanase activity (Fig. 2, band a) and was matched by MS analysis to adenosine diphosphate glucose pyrophosphatase (AGPPase) and germin-like proteins. One of the main roles of AGPPase is hydrolysis of ADPglucose, which is the universal starch precursor, yielding glucose-1-phosphate and AMP and therefore this enzyme determines the net rate of starch synthesis (Rodrígues-López et al., 2000). Interestingly, it was previously found that sucrose levels were slightly increased in Stakado after inoculation with S. tritici, i.e. at 7 dai and 11–15 dai (Shetty et al., 2007). This sugar release essentially correlated with the peaks in AGPPase activity. AGPPases are generally considered to be located in the plastids and has been reported from wheat leaves before, although some have been reported outside the plastids in barley endosperm tissue (Rodrígues-López et al., 2000). However, Rodrígues-López et al. (2001) found that two isoforms of AGPPase were oligomers of the germin-like protein HvGLP1. The protein identified here showed 92% identity to the proteins described by Rodrígues-López et al. (2001), one of which was found to be soluble. In accordance with this, the protein was found to be present in the apoplast after removal of the signal peptide, indicating its secreted nature. Rodrígues-López et al. (2001) furthermore found that neither of the oligomers had oxalate oxidase or superoxide dismutase activity, confirming results by Vallelian-Bindschedler et al. (1998). On the other hand, they suggested that AGPPases may regulate the biosynthesis of cell wall polysaccharides, glycoproteins, and glycolipids by controlling the level of nucleotide sugars. In accordance with this, it has previously been shown that the gene HvGLP1 is constitutively expressed in barley infected by Blumeria graminis f.sp. hordei, but that expression declined under infection, causing the disappearance of the protein at the same time as cell wall reinforcement occurred (Vallelian-Bindschedler et al., 1998; Schweizer et al., 1999) and Schweizer et al. (1999) suggested that HvGLP1 may play a role in stressed leaves, for example, by serving as the substrate for cell-wall reinforcement. Previously, Segarra et al. (2003) reported a germin-like protein with SOD-activity and serine protease-inhibiting activity from the apoplast of wheat infected by S. tritici. However, the relationship to HvGLP1 could not be confirmed by Zimmermann et al. (2006), who did not observe any SOD-activity. Likewise, it has previously been found that even though H2O2 accumulated as a defence response against S. tritici in cv. Stakado (Shetty et al., 2003), there was no SOD activity (NP Shetty et al., unpublished results). Increased cell wall reinforcement was observed in the resistant cv. Stakado during the late stages of the interaction with S. tritici in the form of deposition of callose and, furthermore, cell wall glycoproteins accumulated in the apoplastic fluid, e.g. extensin (Smallwood et al., 1995) and arabinogalactan proteins as well as different types of pectin fragments from the primary plant cell wall matrix. Vallelian-Bindschedler et al. (1998) reported that the germin-like protein HvGLP1 disappeared from the apoplastic fluid of barley infected by the pathogen B. graminis f.sp. hordei, but not after inoculation with the non-host pathogen B. graminis f.sp. tritici, thus indicating that the insolubilization reflected infection-related stress rather than resistance. In the wheat–S. tritici interaction the reverse situation was observed, with a correlation between resistance to infection and glycoproteins in the apoplastic fluid. Thus, the levels of glycoproteins, including extensins, in the apoplastic fluid was low in the susceptible Sevin and high in the resistant Stakado. Apparently, the glycoproteins were released from the cell walls in the leaves. Extensins are hydroxyproline-rich glycoproteins and two major roles for extensins have been suggested (Wei and Shirsat, 2006). Thus, they may provide strengthening of the cell wall by cross-linking and/or anchorage for lignification in order to form a physical barrier to pathogen ingress. In addition, extensins may directly agglutinate around bacteria to prevent further proliferation. Since S. tritici lives in the apoplast and extensins were observed in the apoplastic fluid, the latter role for extensin could be envisaged also to play a role in resistance against this fungal pathogen. Cross-linking of extensins also requires an oxidative burst (Wei and Shirsat, 2006) and this has also been observed in the wheat–S. tritici interaction (Shetty et al., 2003). It was not possible to detect lignin and it was also found that gene expression of two key enzymes in the phenylpropanoid pathway, PAL and CHS, were not elevated after inoculation with S. tritici. Lack of lignification is in agreement with previous studies (Cohen and Eyal, 1993; Kema et al., 1996). Only Ride (1975) reported lignification when wounded wheat leaves were inoculated with S. tritici. Thus, wounding probably potentiated lignin production which accumulated in the presence of the pathogen. The accumulation of callose is often used as a marker for PAMP-elicited defence responses (Kim et al., 2005) and it was found that callose accumulated earlier and to a higher degree in Stakado than in Sevin. The accumulation in Stakado increased dramatically by 5 dai, coinciding with penetration of S. tritici (Shetty et al., 2003). From 7 dai, accumulation was also seen in the substomatal cavities and the mesophyll following the initial growth of the pathogen. In Sevin, only faint callose accumulation was seen and only from 7 dai. Even though Cohen and Eyal (1993) reported callose accumulation in stomata, they did not consider it important for resistance. However, callose has been shown to be deposited on the inner side of the cell walls in response to invasion by micro-organisms and to restrict their growth (Parker et al., 1993). Since S. tritici penetrates through stomata and then grows between the mesophyll cells without penetrating them (Cohen and Eyal, 1993; Kema et al., 1996; Shetty et al., 2003), the accumulation of callose and cell wall glycoproteins from the primary plant cell wall matrix in the apoplastic fluid in response to pathogen growth in cv. Stakado could be envisaged to prevent nutrient and water transfer to the pathogen. In addition, the cell wall reinforcement could offer protection against cell wall-degrading enzymes or toxins produced by the pathogen (Wei and Shirat, 2006). This would help to inhibit the pathogen from spreading in the mesophyll since it derives its nutrients from the host apoplast (Rohel et al., 2001; Keon et al., 2007) in accordance with its hemibiotrophic nature (Parbery, 1996; Rohel et al., 2001; Shetty et al., 2003, 2007). Cell wall-degrading enzymes have been reported from in planta libraries (Kema et al., 2008) and liquid cultures of S. tritici (Douaiher et al., 2007), whereas the existence of toxins have only recently been verified (NP Shetty et al., unpublished results) although the possibility of toxins has been suggested previously (Kema et al., 1996; Shetty et al., 2003, 2007). As a final point, it was tested whether infection by S. tritici could release β-glucan fragments from the pathogen cell walls trough the action of β-1,3-glucanase and whether these β-glucan fragments could act as elicitors, signalling the plant to induce further defence responses. For this, β-glucan was isolated from the pathogen and the susceptible cv. Sevin was treated. First, it was necessary to ascertain that the β-1,3-glucanase released from the plants could cleave the fungal cell walls to release elicitor active molecules, which could act as PAMPs. Infiltration and spraying of purified β-glucans from fungal cell walls 1 dai before inoculation resulted in delayed symptom expression in cv. Sevin, thus efficiently protecting this otherwise susceptible cultivar. In order to verify that wheat was able to degrade the fungal cell walls, a β-1,3-glucanase assay was conducted using apoplastic fluid from cv. Stakado as the enzyme source and purified β-glucans from S. tritici cell walls as the substrate. In plants inoculated with the pathogen, β-1,3-glucanase activity was high and able to degrade the fungal cell walls. Interestingly, the temporal profile of, β-1,3-glucanase activity in the apoplastic fluid in general (Fig. 1B) was quite different from the activity elicited after treatment of purified β-glucan from cell walls (Fig. 7). This is probably due to the secretion of different isoforms during the infection (targeted specifically against the pathogen after penetration) and differences in substrate specificity for laminarin versus fungal β-glucan. In cv. Sevin, there was no up-regulation of the PR-proteins or structural defence responses during the early stages of the interaction (Tables 2, 3). Probably, β-1,3-glucanase isoforms required to cleave the fungal cell walls were not elicited, meaning that release of β-glucan fragments did not take place. To test if defence responses could be elicited in cv. Sevin, plants were treated with purified β-glucan fragments one day before inoculation. There was no difference in fungal penetration (data not shown). However, transcript accumulation of β-1,3-glucanase and chitinase increased and so did callose accumulation, especially in the stomata, indicating that changes in host defence occurred after penetration. Thus, in cv. Sevin, the pathogen is apparently not recognized since PR-protein accumulation is virtually absent, meaning that the fungal cell walls cannot be cleaved and that elicitors and further defence responses are therefore absent. This suggests that when S. tritici establishes contact with a leaf surface of a resistant cultivar, H2O2 accumulates (Shetty et al., 2003), and this could be the signal for the accumulation of PR-proteins as also shown by Ray et al. (2003). When the pathogen penetrates through the stomata, the PR-proteins already present hydrolyse the fungal cell walls, releasing fungal cell wall fragments acting as PAMPs, which induce structural defence responses such as callose, as well as the accumulation of cell wall glycoproteins in the apoplastic fluid (Table 2), to inhibit pathogen growth and reduce the availability of nutrients and water and to protect against fungal cell wall-degrading enzymes and toxins. 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This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.5/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details) © 2009 The Author(s).

Bot, Jacques Le; Bénard, Camille; Robin, Christophe; Bourgaud, Frédéric; Adamowicz, Stéphane

doi: 10.1093/jxb/erp271pmid: 19741002

Abstract Plants allocate internal resources to fulfil essential, yet possibly conflicting, demands such as defence or growth, as hypothesized by the ‘growth–differentiation balance theory’ (GDB). This trade-off was examined in young tomato plants grown for 25 d using the nutrient film technique with seven nitrate concentrations ([NO3]). The modification of primary (growth-related: organic acids, carbohydrates) and secondary (defence-related: phenolics) metabolite concentrations in leaves was assessed. Then a simple model was devised to simulate the trade-off between growth and secondary metabolism in response to N nutrition. N affected growth and metabolite concentrations in the leaves. Dry biomass, leaf area, and concentrations of nitrate and organic acid (malic, citric) increased with rising [NO3], up to a threshold, above which they remained constant. Starch, sucrose, and organic N concentrations were invariant with [NO3]. Glucose, fructose, and phenolic (chlorogenic acid, rutin, and kaempferol-rutinoside) concentrations were highest at lowest [NO3]. They declined progressively with rising [NO3] until a threshold, above which they remained constant. Model predictions are in phase with experimental phenolic concentration data although the simulated metabolic rates differ from the GDBH proposals depicted in the literature. From the model output it is shown that a careful definition of the C reserve compounds is required. Growth–differentiation balance hypothesis (GDBH), leaf composition, major phenolics (chlorogenic acid, rutin and kaempferol-rutinoside), model, nitrate limitation, primary C compounds, Solanum lycopersicum L. (formerlyly Lycopersicon esculentum Mill., tomato) Introduction During evolution, plants have developed adaptive defensive responses against stresses such as grazing insects, microorganisms, or abiotic factors such as UV radiation. Among these, the mechanism of elicitation involves the stimulation of specific metabolic pathways that trigger syntheses and accumulation of a myriad of defence-related metabolites in plant tissues. Growing plants are, therefore, continuously facing a dilemma regarding the partitioning of their available C resources. If priority is given to the plant growth processes, the availability of C resources (and other nutrients) may become limiting for plant defence-related processes, and vice versa. So far, four main plant defence hypotheses have been put forward to explain patterns and variations in the concentration of carbon-based secondary compounds in plant tissues, according to availability of resources: the carbon–nutrient balance hypothesis (CNBH; Bryant et al., 1983; Tuomi, 1992), the optimal defence theory (ODT; McKey, 1974, 1979), the protein competition model (PCM; Jones and Hartley, 1999), and the growth–differentiation balance hypothesis (GDBH; Loomis, 1932; Herms and Mattson, 1992; Herms, 2002). These four hypotheses form a conceptual framework for the functional ecology of plant defence. They suggest that plants continuously make an effective use of costly versus beneficial investments towards defence versus growth processes, the trade-off being mainly conditioned by resource availability, such as N to which this paper is restricted. Many research papers (Herms and Mattson, 1992; Stamp, 2004; Mattson et al., 2005; Glynn et al., 2007) have focused on the GDBH. Herms and Mattson (1992) introduced a simple diagram (Fig. 1) to depict non-linear effects of nutrient availability on secondary metabolism. This representation is widely adopted (Stamp, 2004; Matyssek et al., 2005; Glynn et al., 2007). As the main determinant of this basic trade-off, the GDBH assumes that a shortage of nutrients produces differential effects on plant growth and photosynthetic rates, the former being more restricted than the latter. Considering growth as the product of leaf area by the rate of net CO2 assimilation, the main effect of N supply on plants is to increase leaf area duration (Gastal and Lemaire, 2002). Therefore, at a rate of N supply low enough to alter leaf area, but not so low as to lower the net CO2 assimilation rate, it follows that extra C accumulates in the plants and it may fuel the rate of secondary metabolism. Figure 1 sketches Herms and Mattson's views about the different effects of nutrient availability on the net CO2 assimilation rate (NAR), the plant relative growth rate (RGR), and the rate of secondary metabolism. The diagram explores two contrasted domains of ‘severe nutrient deficiency’ and ‘agricultural practices’, respectively. The former (zone 1) is rarely described in research papers on agricultural crops, but it is tackled by ecological studies on poor habitats (Lambers and Poorter, 1992; Tjoelker et al., 2005). Therein, nutrient availability limits NAR more than RGR and allows extra C substrates to flow into secondary metabolism. The latter domain, however, comprises most scenarios occurring with cropping systems. Therein, nutrient availability sustains maximum NAR due to regular fertilization practices, although these may not be sufficient to maintain maximum growth if there are still insufficient nutrients to give maximum leaf area. Therefore, depending on fertilization efficiency, this domain comprises two contrasted zones, in which nutrition is limiting (zone 2) or non-limiting (zone 3) for RGR. At the transition between zones 2 and 3, plant nutrient status is considered ‘optimal’, also defined elsewhere as critical (Lemaire, 1997) with respect to total plant dry biomass production. It follows that upon increase in nutrient availability, the rate of secondary metabolism is meant to increase over zone 1 and to decrease thereafter, as more of the assimilated C goes into growth processes. Fig. 1. Open in new tabDownload slide Hypothetical response of net assimilation rate (NAR), relative growth rate (RGR), and relative rate of secondary metabolism to N availability, after Herms and Mattson (1992). Fig. 1. Open in new tabDownload slide Hypothetical response of net assimilation rate (NAR), relative growth rate (RGR), and relative rate of secondary metabolism to N availability, after Herms and Mattson (1992). In the case of nitrogen (N), results from the literature either corroborate or contradict this hypothesis. Slow growing N-limited plants generally exhibit elevated concentrations of C-based secondary metabolites (Herms and Mattson, 1992; Stout et al., 1998; Hoffland et al., 2000; Stewart et al., 2001). As a result of metabolic competition between growth-related and defence-related metabolism, reduced spring wheat growth was observed after activation of defence mechanisms (Heil et al., 2000). This series of results indicates some kind of trade-off that corroborates the GDBH. Results are not always in accordance with the GDBH, especially because secondary metabolites form a wide diversity of compounds. As an example, Koricheva (1998) carried out a meta-analysis on literature data of woody plants subjected to various environmental manipulations (N or P fertilization, shading, CO2 enrichment, drought stress, ozone exposure) and noticed that phenolics conformed to the GDBH predictions while terpenes and tannins did not. This may support the view that none of the current theories on resource limitations to secondary metabolism can account for all observed responses, emphasizing the need for a more detailed physiological, and biochemically informed, model (Lou and Baldwin, 2004). Few attempts have yet tried to model the growth–defence trade-off in plants. Most trials concerned trees and herbs (Gayler et al., 2004; see Matyssek et al., 2005; Gayler et al., 2008) and, to our knowledge, annual crops were scarcely studied. At least three publications examined how resource availability (light, water, and N) influenced allocation to defence and growth in tomato (Wilkens et al., 1996a, b; Stamp et al., 2004). Their results were compatible with the GDBH predictions, although the authors estimated that the low number of treatments (only two levels of resource availability) hindered the rigorous testing of the theory. There is a need, therefore, to test experimentally the relationships between C acquisition, growth, and secondary metabolism in tomato along a wide-ranging gradient of resource availability, N for example. However, since the direct testing of the GDBH appears impossible (Stamp, 2004) there is also a need to test the theory through modelling and to confront the experimental patterns with output simulations. The issues for a more comprehensive understanding of these relationships are the control of plant N nutrition in relation to the promotion of natural plant defence and crop production. The first objective of this work is to assess the patterns of the concentration of primary and secondary metabolites in leaves of young hydroponically grown tomatoes, in response to a wide range of nitrate concentrations in the nutrient solution. The second objective is to model these patterns and revisit the GDB hypothesis in the light of the experimental data, for a parameterization of the GDBH components (i.e. rates of C acquisition, growth, and secondary metabolism). The present strategy was (i) to acquire experimental data on young tomato plants (Solanum lycopersicum L.) raised during their exponential growth phase, at contrasting levels of N availability, ranging from limiting to non-limiting for growth (i.e. zones 2–3 in Fig. 1); and (ii) to build up from these data a simple growth model based on the assumptions of Stamp (2004). The specific growth conditions are particularly important as young plants avoid self-shading and metabolic changes due to plant ontogeny. N limitation was assessed by dry biomass accumulation and composition of N resources (i.e. NO3 and organic N) in the leaves. Secondary metabolism was characterized by the quantification of phenolic compounds, particularly chlorogenic acid, rutin, and kaempferol-rutinoside, reported to be the main secondary metabolites in tomato leaves. Chlorogenic acid is a hydroxycinnamic derivative that is quantitatively important in the Solanaceae (Clifford, 1999), and rutin and kaempferol-rutinoside are two examples of plant defence-mediated glycosylated flavonoids (Treutter, 2006). Several primary C compounds were also quantified: organic acids and non-structural carbohydrates such as glucose, fructose, sucrose, and starch. Model overview The model comprises three pools of different biochemical compositions. To compare them, their sizes are expressed in equivalent C (i.e. mmol of C per plant), not in grams. These pools are the non-structural carbon resources (Wc, comprising soluble and insoluble carbohydrates such as sucrose, fructose, glucose, and starch) from which primary and secondary metabolites are built, the C-based secondary compounds (W2), and the remainder, named hereafter metabolic (Wm) as it contains all the metabolic machinery. Total plant mass (Wt, mmol of C per plant) aggregates these pools: (1) The respective concentration of these pools (i.e. Cc, Cm, and C2, mol ratios, dimensionless) is given by: (2) The rate at which Wc accumulates is the net balance between C gain, loss, and allocation to metabolic and secondary stores: (3) where P and R (both in mmol of C per plant d−1) are gross photosynthesis and respiration, respectively. P is the product of the gross assimilation rate (GAR, mmol C cm−2 d−1) and leaf area (A, cm2 per plant): (4) Sinclair (1991) reviewed many studies showing a good correlation between GAR and leaf N concentration ([N], mol N mol−1 C). The general pattern is a linear increase at lower [N], followed by a curvilinear response approaching a maximum rate (Pmax, mmol C cm−2 d−1) at higher [N] admittedly described by the threshold model of Sinclair and Horie (1989): (5) where S is the initial steepness and GAR=0 when [N] equals [N0]. In the absence of a specific model, it is considered that leaf area develops in proportion (α, cm2 mmol−1 C) to the size of the metabolic pool during the exponential growth phase: (6) Respiration (R) comprises two components, growth (Rg) and maintenance (Rm), respectively (Thornley, 1970; McCree, 1983; Thornley and Johnson, 1990): (7) Growth respiration (Rg) is the main component arising from the new synthesis of plant material from Wc. Therefore, it is proportional to the growth rate of both metabolic and secondary pools according to a conversion efficiency or yield (Yg, dimensionless) of the growth process (Thornley and Johnson, 1990). To keep the model simple, Yg was considered equal for both pools: (8) Maintenance respiration (Rm) powers the re-synthesis of degraded material (mainly organic N turnover) and maintenance of concentration gradients in the plants. McCree (1983) proposed to relate Rm linearly to plant N content via a maintenance coefficient (m, mol C mol−1 N per day): (9) The rate of C allocation (mmol of C per plant d−1) to the metabolic pool depends on the size of this pool (i.e. allowing for exponential growth) and on the availability of non-structural carbon resources according to a saturable function (Thornley and Johnson, 1990). Instead, its response to [N] is usually considered linear (Ingestad and Ågren, 1992) as shown for tomato (Martinez et al., 2005), thus: (10) with Vk in mol C mol−1 N d−1 and K1 dimensionless. The rate of C allocation (mmol C per plant d−1) to the C-based secondary compounds pool depends on the availability of non-structural carbon resources and on the size of the metabolic pool since this store encompasses all the metabolic machinery required in the pathways. This rate may thus be modelled as a saturable function such as: (11) with K2 dimensionless and V2max (maximum rate of synthesis) in d−1. The overall plant RGR (d−1) and the RGRs of the metabolic (Mm, d−1) and secondary (M2, d−1) pools are calculated as: (12) Numerical integration of the differential equations (i.e. Equations 3, 10, and 11) relates biomass accumulation in the three pools to [N]. In order to relate it to the concentration of N in the nutrient medium, as proposed in Fig. 1, it is necessary to introduce an N uptake submodel. For this purpose, the model of Cárdenas-Navarro et al. (1999), developed and parameterized on young tomato plants, was used. The uptake rate (I, mol N mol C−1 d−1) responds to NO3 concentration (ns, mol m−3) in the nutrient solution following saturable Michaelis–Menten kinetics. Furthermore, it responds negatively to plant NO3 concentration, but their model was simplified by replacing plant NO3 by [N] as follows: (13) with Ks in mol m−3, Imax being the maximum influx rate when [N] tends to zero and ns tends to infinity. I=0 when [N]=[N]max. Assuming the N uptake rate is proportional to the importance of the metabolic machinery, the plant nitrogen content (QN, mol of N per plant) increases with time at the following rate: (14) As QN=[N]×Wt, it follows that: (15) thus, combining Equations 14 and 15 yields : (16) which, according to Equations 2 and 12, can be written: (17) Numerical solving of this latter differential allows the plant N status to be related to N nutrition and to growth. Materials and methods Experimental set-up The plants were grown in Avignon (France) on seven independent recycling NFT (nutrient film technique) systems randomly arranged in a heated greenhouse equipped with mist spray (see a comprehensive description in Adamowicz and Le Bot, 2008). On each system, 42 plants were raised with a nutrient solution of constant nitrate concentration ([NO3]) flowing at the rate of ∼0.6 l min−1. [NO3] in treatments were 0.050, 0.125, 0.2, 0.3, 3.0, 7.0, and 15.0 mol m−3, and their volumes of solution were, respectively, 1, 1, 0.7, 0.7, 0.3, 0.3, and 0.3 m3 per treatment. Higher volumes were used with lower concentrations to help limit [NO3] drifts due to plant ion uptake. The drifts in solution ionic composition were monitored from manual sampling and [NO3] and pH measurements, with an increasing frequency as growth proceeded, up to twice a day. Based upon analyses, NO3 salts and H2SO4 were added to compensate for drifts. At the end of the experiment it was verified that the cationic concentrations did not deviate significantly from pre-set values. The greenhouse was whitewashed to reduce incident light intensity on the crops and to make air temperature control easier. Routine climatic measurements included plant photosynthetic flux density (PPFD) at crop level (QS sensors, Delta-T Devices, Cambridge, UK) and temperature (thermistance probes ±0.1 °C, TJI18-44043-1/4-2, Newport Omega, Stamford, CA, USA) of air under cover and of each of the nutrient solutions. Plant material and growth conditions The tomato plants (Solanum lycopersicum L. cv Rondello F1, De Ruiter seeds, Bleiswijk-Holland) were sown directly in the NFT system (9 September 2000). The greenhouse was heated at 18 °C and aerated at 25 °C during germination and emergence (9 d), then aerated at 20 °C. Relative humidity was regulated at ∼55%. To avoid any mechanical stress, plants were never handled until harvest. All nutrient solutions, except the 15 mol m−3 nitrate solution, were devised to make the sum (NO3– + SO42–)=12 eq m−3, inferring constant concentrations of other ionic species in all treatments. Nutrient solutions were prepared from deionized water and pure salts at the following concentrations (eq m−3): K 3; Ca 7; Mg 3; H2PO4 1; trace elements were provided as Kanieltra (Hydro Azote, France) formula 6 Fe (0.1 l m−3) and EDTA-Fe (43 μM). Inevitably the treatment at 15 mol m−3 NO3 contained more salts (eq m−3): SO4 5; K 4; Ca 12.2; Mg 4.8; H2PO4 1. Nonetheless, all recipes shared analogous chemical activity ratios for major cationic species. Solution pH was 5.5 and electrical conductivity (EC, mS cm−1) ranged from 1.2–1.4 to 2.1 depending on treatment. Harvest and sample preparation The experiment lasted for 28 d, during which five harvests were taken 14, 18, 21, 25, and 28 days after sowing (das). Plants were harvested at the end of the night period (6 am). Light has pronounced effects on plant metabolite concentrations (Fritz et al., 2006) but it is observed that these changes over the diel cycle are of least influence at the end of the night period. Thus, under natural climate, harvesting plants at the end of the night period minimizes the variability due to PPFD changes during the previous day. Plants were sampled in organ classes (leaves, stems, roots), which were analysed separately for non-structural compounds. All leaf samples collected 14, 18, 21, and 25 das were analysed for phenolics but, due to low biomass, only a few stem and root fractions sampled 25 das were examined. In this paper, only leaf data (≥60% of total plant dry biomass) obtained at the fourth harvest (25 das) from eight plants per treatment are presented and used for the model. The plants were sampled at random and at least one plant was selected per growing tray. The other plants remained on the set-up in isolated growth conditions with respect to light interception. All plants were stored in a dark cold room (15 °C, 1–3 h) before their measurements. Leaf area was measured on an area meter (LI-3000A with LI-3050A belt conveyor, LI-COR, Lincoln, NE, USA) and fresh biomass was weighed on a precision balance (model AE 260, sensitivity 0.1 mg, Mettler, Greifensee, Switzerland). All samples were frozen in liquid nitrogen and kept at –20 °C. For the harvest reported herein, the entire process concerned 56 plants and lasted for ∼3 h. Samples were freeze-dried (Genesis 25 ES, Virtis Company, Gardiner, NY, USA), weighed (model AE 163, sensitivity 0.01 mg, Mettler), and ground to a fine powder (model MM200, Retsch, Haan, Germany) in grinding jars, which were pre-cooled in liquid nitrogen. Dry powders were kept at –20 °C until analysed. Chemical analyses Solution nitrate concentration was measured by UV spectrometry using the method developed by Vercambre and Adamowicz (1996). Plant tissue nitrate was determined on water extracts of dried powders using an autoanalyser (AQUATEC 5500, Tecator, Höganäs, Sweden) by colorimetry of nitrite after reduction by cadmium. Total sample N and C were determined according to the Dumas method (elemental analyser NA 1500, Carlo Erba, Milano, Italy). Organic N was computed as the difference between total and nitrate N. Starch was extracted and determined according to the procedure of Gomez et al. (2003). Soluble sugars and carboxylic acids were extracted and measured by HPLC following the method described by Gomez et al. (2002) and Wu et al. (2002). In the spectrum of acids, only malic and citric are reported here, but low concentrations of fumaric (<0.2% DW) and shikimic (<0.05% DW) were also detected in the extracts. Phenolics were extracted using the method of Fleuriet (1976) modified as follows. All steps were carried out at 4 °C either in a cold chamber or on ice. Dry powder (50 mg) was extracted three times with 2 ml of 70% aqueous ethanol. Taxifolin solution (50 μl of a stock at 2 mg ml−1 methanol) was added as an internal standard. Each time the mixture was blended for 1 min and homogenized for 15 min. After centrifugation, supernatants were pooled to constitute the raw extract. The extract (total volume: 6 ml) was evaporated to dryness under vacuum. The residue was dissolved in 0.4 ml of methanol and filtered through a 0.45 μm filter (Minisart RC 4, Sartorius) prior to injection (20 μl) into the HPLC. The analytical conditions were those described by Gautier et al. (2008). All compounds were correctly separated (not shown), allowing the quantification of rutin, chlorogenic acid, and kaempferol-rutinoside from peak area calibrated against a standard curve. Rutin and chlorogenic acid standards were purchased from Sigma (Saint Quentin-Fallavier, France), and kaempferol-rutinoside and taxifolin from Extrasynthèse (Lyon, France). Data processing Changes in leaf growth or composition (y) in response to nitrogen nutrition (x) were analysed using a broken stick procedure whereby data were fitted with two successive linear regressions y=a1×x+b1 and y=a2×x+b2, joining at dose (xbp) called the ‘breakpoint’. At this point y=a1×xbp+b1=a2×xbp+b2 and, as a result, it follows that: (18) In order to determine the parameters of the regressions, a statistical analysis was carried out with the software SYSTAT (v. 5.1 for the Macintosh, SYSTAT Inc., Evanston, IL, USA) using the piecewise linear regression procedure with an unknown breakpoint, with the general formula: (19) where y is the explained variable (either growth or leaf concentration), x is solution [NO3], b1 the constant, and a1 and a2 the slopes before and after the breakpoint (xbp), respectively. Parameters were found by optimization. The statistical significance of the slopes was assessed, especially above xbp where the asymptotic nature of the response is supposed to intervene. Moreover, if a1 and a2 did not prove significantly different, a single linear regression was calculated over the entire range of [NO3] treatments. In order to represent correctly the 300-fold changes in [NO3] on a linear scale, the data were plotted in a broken axis chart (DeltaGraph v 5.7.5 for the Macintosh, SPSS Inc. & Red Rock Software, Inc., Chicago, IL, USA), which skipped the value axis portion where 0.4 < [NO3] <3 mol m−3 so as to scale specifically the two regions that varied clearly from each other. Numerical solving of the model was performed with ExtendSim software (v.7, Imagine That, Inc., San Jose, CA, USA). Results Experimental data At 25 das, leaves, stems, and roots accounted for 60, 30, and 10% of plant DW, respectively. It was found (not shown) that the chemical composition of both shoot pools shared the same response patterns to N. Leaf polyphenol concentrations were higher than in stems and roots, making leaves a suitable indicator of the whole plant. Leaf dry biomass (DW, g per plant) and leaf area (LA, cm2 per plant) responded to NO3 availability and, according to the piecewise regression analysis, the breakpoint occurred in the region of 0.3 mol m−3 NO3 in solution (Table 1, Fig. 2). At the most, low NO3 treatment decreased DW and LA by ∼30%. Above the breakpoint, DW was not significantly enhanced by NO3 availability, although LA was slightly augmented by increasing N nutrition. This clear-cut phase transition segregates the limiting from the adequate domain of N availability for growth. In the former, growth was ∼60 times more responsive to NO3 than in the latter, as shown by the calculation of the slope ratio (Table 1). Leaf NO3 concentration was markedly affected by the treatment (Table 1, Fig. 3 circles, dotted line). Under low N, it increased sharply with plant N nutrition and above the breakpoint it remained constant. Reduced N concentration was not significantly affected by the treatment (Table 1, Fig. 3 squares, thin line). Overall, it resulted that increasing NO3 availability in the nutrient solution increased total N leaf concentration slightly, but significantly (Table 1, Fig. 3, triangles, thick line). In the adequate N treatments, leaf N NO3 accounted for ∼10% of the total N store. Table 1. Parameters of piecewise linear regression analyses with an unknown breakpoint (xbp) for leaf growth (DW, dry weight; LA, leaf area) and chemical concentrations ([NO3], nitrate; [Nt], total N; [Nr], reduced N; OA, organic acids; KR, kaempferol-rutinoside) DW (g) LA (cm2) [NO3] (% in DW) [Nt] (% in DW) [Nr] (% in DW) Glucose (% in DW) Fructose (% in DW) Sucrose (% in DW) b1 0.196* 111.2* −0.037 5.761* 5.365* 1.449* 3.489* 1.158* a1 0.247* 156.1* 11.421* 0.048* 0.028 −5.975* −9.823* 0 xbp 0.308* 0.3* 0.214* – – 0.156* 0.186* – b2 0.271* 161.5* 2.401* 0.514* 1.657* a2 0.004 2.8* 0.029 0.017* 0.025* Ratio a1/a2 62 57 394 351 393 DW (g) LA (cm2) [NO3] (% in DW) [Nt] (% in DW) [Nr] (% in DW) Glucose (% in DW) Fructose (% in DW) Sucrose (% in DW) b1 0.196* 111.2* −0.037 5.761* 5.365* 1.449* 3.489* 1.158* a1 0.247* 156.1* 11.421* 0.048* 0.028 −5.975* −9.823* 0 xbp 0.308* 0.3* 0.214* – – 0.156* 0.186* – b2 0.271* 161.5* 2.401* 0.514* 1.657* a2 0.004 2.8* 0.029 0.017* 0.025* Ratio a1/a2 62 57 394 351 393 Starch (% in DW) Citric (% in DW) Malic (% in DW) Total OA (% in DW) Chlorogenic acid (‰ in DW) Rutin (‰ in DW) KR (‰ in DW) b1 5.919* −0.103 0.469 -0.554 3.142* 0.613* 0.472* a1 −0.188 5.608* 6.951* 12.653* −4.726* −1.003* −0.543* xbp – 0.444* 0.699* 0.584* 0.302* 0.367* 0.238* b2 2.342* 4.286* 6.688* 1.723* 0.243* 0.342* a2 0.102 0.148* 0.253* −0.028 0.005 0.002 Ratio a1/a2 55 47 50 168 200 270 Starch (% in DW) Citric (% in DW) Malic (% in DW) Total OA (% in DW) Chlorogenic acid (‰ in DW) Rutin (‰ in DW) KR (‰ in DW) b1 5.919* −0.103 0.469 -0.554 3.142* 0.613* 0.472* a1 −0.188 5.608* 6.951* 12.653* −4.726* −1.003* −0.543* xbp – 0.444* 0.699* 0.584* 0.302* 0.367* 0.238* b2 2.342* 4.286* 6.688* 1.723* 0.243* 0.342* a2 0.102 0.148* 0.253* −0.028 0.005 0.002 Ratio a1/a2 55 47 50 168 200 270 xbp is the solution [NO3] dividing N-limiting from adequate nutrition domains. b1 and a1 are the constant and the slope of the linear regression before xbp, respectively. b2 and a2 are the constant and the slope of the linear regression after xbp. The slope ratio (a1/a2) assesses the intensity of leaf response change between the two domains. An asterisk indicates that the parameter is significant (P=0.05). When xbp is not significant (–) the regression parameters are those calculated over the entire range of N supply. Open in new tab Table 1. Parameters of piecewise linear regression analyses with an unknown breakpoint (xbp) for leaf growth (DW, dry weight; LA, leaf area) and chemical concentrations ([NO3], nitrate; [Nt], total N; [Nr], reduced N; OA, organic acids; KR, kaempferol-rutinoside) DW (g) LA (cm2) [NO3] (% in DW) [Nt] (% in DW) [Nr] (% in DW) Glucose (% in DW) Fructose (% in DW) Sucrose (% in DW) b1 0.196* 111.2* −0.037 5.761* 5.365* 1.449* 3.489* 1.158* a1 0.247* 156.1* 11.421* 0.048* 0.028 −5.975* −9.823* 0 xbp 0.308* 0.3* 0.214* – – 0.156* 0.186* – b2 0.271* 161.5* 2.401* 0.514* 1.657* a2 0.004 2.8* 0.029 0.017* 0.025* Ratio a1/a2 62 57 394 351 393 DW (g) LA (cm2) [NO3] (% in DW) [Nt] (% in DW) [Nr] (% in DW) Glucose (% in DW) Fructose (% in DW) Sucrose (% in DW) b1 0.196* 111.2* −0.037 5.761* 5.365* 1.449* 3.489* 1.158* a1 0.247* 156.1* 11.421* 0.048* 0.028 −5.975* −9.823* 0 xbp 0.308* 0.3* 0.214* – – 0.156* 0.186* – b2 0.271* 161.5* 2.401* 0.514* 1.657* a2 0.004 2.8* 0.029 0.017* 0.025* Ratio a1/a2 62 57 394 351 393 Starch (% in DW) Citric (% in DW) Malic (% in DW) Total OA (% in DW) Chlorogenic acid (‰ in DW) Rutin (‰ in DW) KR (‰ in DW) b1 5.919* −0.103 0.469 -0.554 3.142* 0.613* 0.472* a1 −0.188 5.608* 6.951* 12.653* −4.726* −1.003* −0.543* xbp – 0.444* 0.699* 0.584* 0.302* 0.367* 0.238* b2 2.342* 4.286* 6.688* 1.723* 0.243* 0.342* a2 0.102 0.148* 0.253* −0.028 0.005 0.002 Ratio a1/a2 55 47 50 168 200 270 Starch (% in DW) Citric (% in DW) Malic (% in DW) Total OA (% in DW) Chlorogenic acid (‰ in DW) Rutin (‰ in DW) KR (‰ in DW) b1 5.919* −0.103 0.469 -0.554 3.142* 0.613* 0.472* a1 −0.188 5.608* 6.951* 12.653* −4.726* −1.003* −0.543* xbp – 0.444* 0.699* 0.584* 0.302* 0.367* 0.238* b2 2.342* 4.286* 6.688* 1.723* 0.243* 0.342* a2 0.102 0.148* 0.253* −0.028 0.005 0.002 Ratio a1/a2 55 47 50 168 200 270 xbp is the solution [NO3] dividing N-limiting from adequate nutrition domains. b1 and a1 are the constant and the slope of the linear regression before xbp, respectively. b2 and a2 are the constant and the slope of the linear regression after xbp. The slope ratio (a1/a2) assesses the intensity of leaf response change between the two domains. An asterisk indicates that the parameter is significant (P=0.05). When xbp is not significant (–) the regression parameters are those calculated over the entire range of N supply. Open in new tab Fig. 2. Open in new tabDownload slide Leaf dry biomass (A, g per plant) and area (B, cm2 per plant) as a function of NO3 concentration in the nutrient solution. Symbols and bars are means and standard errors calculated on eight replicate plants per N treatment. The dashed lines are piecewise linear regressions (parameters in Table 1). Note the x-axis break between 0.4 and 3 mol NO3 m−3. Fig. 2. Open in new tabDownload slide Leaf dry biomass (A, g per plant) and area (B, cm2 per plant) as a function of NO3 concentration in the nutrient solution. Symbols and bars are means and standard errors calculated on eight replicate plants per N treatment. The dashed lines are piecewise linear regressions (parameters in Table 1). Note the x-axis break between 0.4 and 3 mol NO3 m−3. Fig. 3. Open in new tabDownload slide Leaf nitrogen concentration ([N], g per 100 g DW) as a function of NO3 concentration in the nutrient solution. The graph displays NO3 N (circles, dotted line), organic N (squares, thin line), and total N (triangles, thick line) concentrations. The lines are piecewise linear regressions (parameters in Table 1). Symbols and bars are means and standard errors. Fig. 3. Open in new tabDownload slide Leaf nitrogen concentration ([N], g per 100 g DW) as a function of NO3 concentration in the nutrient solution. The graph displays NO3 N (circles, dotted line), organic N (squares, thin line), and total N (triangles, thick line) concentrations. The lines are piecewise linear regressions (parameters in Table 1). Symbols and bars are means and standard errors. Leaf starch concentration was not significantly affected by N supply, although starch tended to accumulate at low N nutrition (Table 1, Fig. 4a). Similarly, leaf sucrose concentration was insensitive to N supply (Table 1, Fig. 4b). Hexose concentration, however, responded to N nutrition (Table 1, Fig. 4b) as both fructose and glucose decreased sharply in the limiting N domain before they increased slightly (but significantly, Table 1) in the non-limiting zone. The responses were far more intensive under N limitation, as assessed by the slope ratio of 350–400 calculated between the two regressions (Table 1). Under non-limiting N, the leaf accumulated three times more fructose than glucose, making total hexose account for 2.4% of leaf dry biomass. In comparison, leaf starch represented up to 4.7% of the leaf dry biomass. Fig. 4. Open in new tabDownload slide Leaf starch (A, g per 100 g DW) and soluble carbohydrate (B, g per 100 g DW) concentrations as a function of NO3 concentration in the nutrient solution. Symbols in (B) denote sucrose (open circles), glucose (filled circles), and fructose (filled squares). The lines are piecewise linear regressions (parameters in Table 1). Symbols and bars are means and standard errors. Fig. 4. Open in new tabDownload slide Leaf starch (A, g per 100 g DW) and soluble carbohydrate (B, g per 100 g DW) concentrations as a function of NO3 concentration in the nutrient solution. Symbols in (B) denote sucrose (open circles), glucose (filled circles), and fructose (filled squares). The lines are piecewise linear regressions (parameters in Table 1). Symbols and bars are means and standard errors. Non-structural leaf organic acid concentration increased markedly with N nutrition (Fig. 5a). Under N limitation, the rate of organic acid accumulation in the leaves was 50 times higher than in the adequate N range (Table 1). Amongst carboxylic acids, malic accumulated almost twice more than citric. Under non-limiting N, free organic acids accounted for 7.3% of the leaf dry biomass, rendering this pool almost as important as non-structural sugars (8.3%) in the leaf C reserve store. Fig. 5. Open in new tabDownload slide Leaf organic acid (A, g per 100 g DW) and phenolic (B, g per 1000 g DW) concentration as a function of NO3 concentration in the nutrient solution. Symbols in (A) denote citric acid (circle, dotted line), malic acid (squares, thin line), and total carboxylic acids (triangles, thick line). Symbols in (B) denote chlorogenic acid (circles, dotted line), rutin (squares, thin line), and kaempferol-rutinoside (triangles, thick line). The lines are piecewise linear regressions (parameters in Table 1). Symbols and bars are means and standard errors. Fig. 5. Open in new tabDownload slide Leaf organic acid (A, g per 100 g DW) and phenolic (B, g per 1000 g DW) concentration as a function of NO3 concentration in the nutrient solution. Symbols in (A) denote citric acid (circle, dotted line), malic acid (squares, thin line), and total carboxylic acids (triangles, thick line). Symbols in (B) denote chlorogenic acid (circles, dotted line), rutin (squares, thin line), and kaempferol-rutinoside (triangles, thick line). The lines are piecewise linear regressions (parameters in Table 1). Symbols and bars are means and standard errors. The phenolic molecules analysed herein were carefully selected amongst dozens of already identified tomato compounds (Moco et al., 2006). The main substances found in the leaf extract of the tomato cv Rondello were chlorogenic acid, rutin, and kaempferol-rutinoside, which are reported to be important secondary compounds in plants (Clifford, 1999; Treutter, 2006). Leaf phenolic concentration responded to N supply (Table 1, Fig. 5b). The pattern was similar for chlorogenic acid, rutin, and kaempferol-rutinoside. Chlorogenic acid was the major leaf phenolic compound. The phenolic concentration decreased rapidly with increasing N availability in the domain of N limitation, but it did not vary significantly under adequate N nutrition. The slope ratio between these regressions ranged from 170 to 270 according to phenolic compounds (Table 1), indicating a more intensive plant response under N limitation. Model simulations Parameterization: The parameters used in the simulations are summarized in Table 2. Their values were either taken from the literature (or derived assuming 40% C in the dry matter, Broadley et al., 2004) or optimized on the present data. Table 2. Model parameters used in the simulations Parameter Units Value in model Equation in use Source Pmax mmol C cm−2 d−1 0.033 5 Optimization [N0] mol N mol−1C 0.00 5 After Chapin et al. (1988) S mol C mol−1N 33.33 5 After Chapin et al. (1988) α cm2 mmol−1 C 25,89 6 Optimization Yg – 0.75 8 Ruget (1981) m mol C mol−1 N d−1 0.38 9 After Gary (1988) Vk mol C mol−1 N d−1 1.31 10 Optimization K1 – 1.7×10−5 10 Optimization V2max d−1 1.60 11 Optimization K2 – 0.501 11 Optimization [N]max mol N mol−1C 0.20 13 Chapin et al. (1988) Ks mol NO3 m−3 0.024 (0–0.08) 0.650 (0.08–7) 13 Cárdenas-Navarro et al. (1999) 17.8 (7–30) Imax mol N mol−1 C d−1 7.85 13 Optimization Parameter Units Value in model Equation in use Source Pmax mmol C cm−2 d−1 0.033 5 Optimization [N0] mol N mol−1C 0.00 5 After Chapin et al. (1988) S mol C mol−1N 33.33 5 After Chapin et al. (1988) α cm2 mmol−1 C 25,89 6 Optimization Yg – 0.75 8 Ruget (1981) m mol C mol−1 N d−1 0.38 9 After Gary (1988) Vk mol C mol−1 N d−1 1.31 10 Optimization K1 – 1.7×10−5 10 Optimization V2max d−1 1.60 11 Optimization K2 – 0.501 11 Optimization [N]max mol N mol−1C 0.20 13 Chapin et al. (1988) Ks mol NO3 m−3 0.024 (0–0.08) 0.650 (0.08–7) 13 Cárdenas-Navarro et al. (1999) 17.8 (7–30) Imax mol N mol−1 C d−1 7.85 13 Optimization Notice that three Ks values are used in the model depending on the [NO3] domain fed to the plants (these domains, expressed in mol NO3 m−3, are given in parentheses, after the Ks values). Open in new tab Table 2. Model parameters used in the simulations Parameter Units Value in model Equation in use Source Pmax mmol C cm−2 d−1 0.033 5 Optimization [N0] mol N mol−1C 0.00 5 After Chapin et al. (1988) S mol C mol−1N 33.33 5 After Chapin et al. (1988) α cm2 mmol−1 C 25,89 6 Optimization Yg – 0.75 8 Ruget (1981) m mol C mol−1 N d−1 0.38 9 After Gary (1988) Vk mol C mol−1 N d−1 1.31 10 Optimization K1 – 1.7×10−5 10 Optimization V2max d−1 1.60 11 Optimization K2 – 0.501 11 Optimization [N]max mol N mol−1C 0.20 13 Chapin et al. (1988) Ks mol NO3 m−3 0.024 (0–0.08) 0.650 (0.08–7) 13 Cárdenas-Navarro et al. (1999) 17.8 (7–30) Imax mol N mol−1 C d−1 7.85 13 Optimization Parameter Units Value in model Equation in use Source Pmax mmol C cm−2 d−1 0.033 5 Optimization [N0] mol N mol−1C 0.00 5 After Chapin et al. (1988) S mol C mol−1N 33.33 5 After Chapin et al. (1988) α cm2 mmol−1 C 25,89 6 Optimization Yg – 0.75 8 Ruget (1981) m mol C mol−1 N d−1 0.38 9 After Gary (1988) Vk mol C mol−1 N d−1 1.31 10 Optimization K1 – 1.7×10−5 10 Optimization V2max d−1 1.60 11 Optimization K2 – 0.501 11 Optimization [N]max mol N mol−1C 0.20 13 Chapin et al. (1988) Ks mol NO3 m−3 0.024 (0–0.08) 0.650 (0.08–7) 13 Cárdenas-Navarro et al. (1999) 17.8 (7–30) Imax mol N mol−1 C d−1 7.85 13 Optimization Notice that three Ks values are used in the model depending on the [NO3] domain fed to the plants (these domains, expressed in mol NO3 m−3, are given in parentheses, after the Ks values). Open in new tab [N0] and S were calculated by applying Equation 5 to the tomato data of Chapin et al. (1988). The maintenance coefficient (m) was derived from the data of Gary (1988) at 21 °C (the mean growth temperature in the present experiment). The model of Cárdenas-Navarro et al. (1999) describes triphasic NO3 influx into tomato with increasing ns, meaning that there are three Ks and Imax values depending on the ns domain. Cárdenas-Navarro et al. (1999) also showed that two values of Imax can be inferred from the knowledge of the three Ks and one Imax. Thus, the values of Ks and the ns domains published by these authors were used, and one Imax was determined by optimization. Optimization followed the procedure described by Wallach et al. (2001) yielding the parameter values that minimized the following goodness-of-fit criterion: (20) Over the seven treatments, (Wt), (A) and ([Cc]) are, respectively, the measured plant weight, leaf area, and non-structural (i.e. carbohydrates) C concentration of treatment (i) while (Wtc), (Ac), and ([Cc]c) are the corresponding value calculated with the model; σW, σA and σCc are the respective standard deviations. Simulation results: Simulations were run with different ns ranging from 0 (low N) to 30 mol NO3 m−3. Plant growth and composition were simulated between 14 (first harvest) and 25 das, and the ultimate simulation data (25 das) were plotted either against ns (Fig. 6a, c) or [N] (Fig. 6b, d). In each run the initial growth values were those of the real experiment. At 14 das, no significant difference was observed between ns treatments; thus the mean Wt (1.32 mmol of C per plant) was used as the initial condition for all treatments. Initially, Wt was arbitrarily allocated 20, 40, and 40% to Wc, Wm, and W2, respectively (Equation 1). Fig. 6. Open in new tabDownload slide Simulated effects of either solution [NO3] (A, C) or plant [N] (B, D) on metabolic rates and C store concentrations. (A, B) Relative rates of growth (RGR, thin line, left axis), secondary metabolism (M2, thick line, left axis), and gross assimilation rate (GAR, dotted line, right axis). (C, D) Concentration of the C stores, where lines are [Cm] (thin), [Cc] (thick), and [C2] (dashed). Simulations were performed from 0 to 30 mol NO3 m−3, but for clarity the plots (A, C) were restricted to 0–1 mol NO3 m−3 as traces were stable beyond this range. Fig. 6. Open in new tabDownload slide Simulated effects of either solution [NO3] (A, C) or plant [N] (B, D) on metabolic rates and C store concentrations. (A, B) Relative rates of growth (RGR, thin line, left axis), secondary metabolism (M2, thick line, left axis), and gross assimilation rate (GAR, dotted line, right axis). (C, D) Concentration of the C stores, where lines are [Cm] (thin), [Cc] (thick), and [C2] (dashed). Simulations were performed from 0 to 30 mol NO3 m−3, but for clarity the plots (A, C) were restricted to 0–1 mol NO3 m−3 as traces were stable beyond this range. As stated in the hypotheses, the rate of C fixation (GAR in the model) is responsive to plant [N] (Fig. 6b), with a sharp increase at low N followed by a low or no increase at high [N], while RGR is nearly linear with [N]. M2 increases non-linearly with [N]. When plotted against ns (Fig. 6a), all the rates (GAR, RGR, and M2) saturate at high NO3 availability. The bumpy patterns are a consequence of the multiphasic uptake model and thus appear solely in Fig 6a. Changes in the concentration of the different plant C pools with N status are plotted in Fig. 6d. [Cc] increases from low to 0.09 mol N mol−1 C and decreases thereafter. [Cm] declines very slightly from low to 0.04 mol N mol−1 C and thereafter increases steadily with [N]. [C2] decreases almost linearly with [N]. When plotted against NO3 availability in the nutrient solution (Fig. 6c), [Cc] shows little response to ns except at very low availability (maximum [Cc] at 7×10−3 mol NO3 m−3). The minimum [Cm] at 8×10−4 mol NO3 m−3 is barely visible but, thereafter, [Cm] responds positively to ns following a multiphasic saturable function. Conversely, [C2] is maximal at low ns and decreases thereafter. All the concentrations ([Cc], [Cm], and [C2]) stabilize at >1 mol NO3 m−3. [C2] relates to [Cc] following a complex pattern (Fig. 7). In the region of strong N limitation (i.e. 0.020 < [N] ≤ 0.089) the relationship is negative. Conversely, at higher N status it becomes positive. Fig. 7. Open in new tabDownload slide Model simulation of [C2] as a function of [Cc] for various N supplies. Tags indicate plant N concentration. Fig. 7. Open in new tabDownload slide Model simulation of [C2] as a function of [Cc] for various N supplies. Tags indicate plant N concentration. Discussion This paper is not a comprehensive test of the GDBH, whose scope is far larger than the particular focus made here on N. The GDB theory has previously been tested on trees (Stamp, 2004; Mattson et al., 2005; Glynn et al., 2007), and dynamic models complementary to GDB have been proposed (Gayler et al., 2004; Matyssek et al., 2005). Similar attempts are lacking for agronomic/annual plants and, due to different plant growth dynamics from perennial species, it is debatable that the GDB theory would properly describe the dynamics of carbon partitioning in the former plants. Growth response to N supply under continuous monitoring Solution [NO3] was tightly controlled and the experiment segregated N limitation (zone 2 of Fig. 1) from adequate N nutrition (zone 3 of Fig. 1) for plant growth (Fig. 2), supporting published statements that large N treatment numbers are required to measure this characteristic (Justes et al., 1994, 1997). The critical solution [NO3] under which growth restriction occurred was low (300 μM) compared with other experimental systems or plant species (range =2.75–12 mM, see Gomez-Lepe and Ulrich, 1974; Gunes et al., 1998; Siddiqi et al., 1998; Tocquin et al., 2003) and with efficient flowing solution culture (FSC), in which solution [NO3] in the range of 10–14 μM can support maximum growth (Parker and Norvell, 1999). This implies that its value depends essentially on the capacity of the growing systems to renew nutrients at the root level. Due to its large number (seven) of N treatments, the present study compares with an experiment on willow trees made by Glynn et al. (2007). It contrasts, however, with many published studies comprising a limited set (two or three) of N treatments (Scheible et al., 1997; Guidi et al., 1998; Stout et al., 1998; Gayler et al., 2004; Urbanczyk-Wochniak and Fernie, 2005). The tomato growth response to N obeys Liebig's law of the minimum and follows a generalized response curve relating plant production to N supply (see Lawlor, 2002). Plant response was analysed according to a broken stick procedure (as in Justes et al., 1994), which fitted the data well, the model accounting for >95% of the observed growth variation. In the leaves, dry biomass accumulation was N limited from 50 μM to 300 μM NO3 in solution (Fig. 2a). The drop in biomass accumulation attained 30% at the most between treatments. This was matched by a similar decrease (34%) of leaf area expansion (Fig. 2b), which inevitably reduced, in proportion, light interception and crop production, inferring that NAR was constant in all treatments. These points and consistent data are thoroughly discussed elsewhere (see Adamowicz and Le Bot, 2008). It must be emphasized that the experimental data did not cover the entire range of N supplies described by Fig. 1 and by the simulation model (Fig. 6). The avoidance of N deficiency per se and the unaltered NAR are corroborated by the lack of variation in leaf organic N concentration, inferring steadiness in operation of the photosynthetic apparatus. Mean leaf N concentration (1.01 g N m−2) appears low in comparison with data on other plants (Sinclair and Horie, 1989), but the present growth data support the general findings that leaf area development is tightly controlled by N availability (Palmer et al., 1996; Lawlor, 2002; Dreccer, 2005). Plant composition in primary and secondary metabolites Many differences in plant composition concerned both primary and secondary metabolites. Under N limitation, leaf [NO3] responded strongly to N nutrition (Fig. 3), but was nearly constant at maximum value under non-limiting N, which fully matches with previous data (Gomez-Lepe and Ulrich, 1974; Scheible et al., 1997; Chen et al., 2004). In contrast, organic N concentration (Fig. 3) was not significantly changed by the treatments, supporting the view that, in tomato, NO3 is the main labile N store responding to environmental changes in N availability (Le Bot et al., 2001). Consistently, this has been used to elaborate practical tools to diagnose plant N status and devise fertilization strategies (Lawlor, 2002; Lemaire et al., 2008). Dissimilar responses occurred in carbohydrate concentrations. Starch, the main leaf C store, did not change concentration over the range of solution [NO3] (Fig. 4a). At first sight this is at odds with literature data reporting on a remarkable negative relationship between N availability and starch level in leaves (Radin and Eidenbock, 1986; Rufty et al., 1988; Paul and Driscoll, 1997; Ball et al., 1998). However, (i) the plants were never N deprived but limited in their supply (Fig. 2a) and (ii) the observation is prone to be influenced by the harvest time (end of the night period). Because starch is strongly depleted overnight (Rufty et al., 1983; Paul and Driscoll, 1997; Ball et al., 1998) to sustain structural growth, all treatments were likely to exhibit comparable low starch concentration at dawn. Similar to starch, the concentration of sucrose (Fig. 4b, circles, solid line) remained constant between N treatments in accordance with other results for tomato (Guidi et al., 1998; Urbanczyk-Wochniak and Fernie, 2005), although the latter reported a different behaviour at low light intensity. Sucrose is a transport sugar whose concentration results from synthesis and export processes. The former is largely controlled by the enzyme SPS (sucrose phosphate synthase), whose activity is perturbed when N deficiency impairs RGR (Foyer et al., 1995). In the present experiment, however, given the unaltered NAR and starch concentration between treatments, unchanged sucrose concentration is likely to reflect similar export activities to sink organs. In contrast, fructose and glucose concentrations were altered at low N nutrition. The pattern was a sharp decrease between 50 μM and ∼200 μM NO3 followed by a slight concentration increase above this value (Fig. 4b). The hexose accumulation rate was 350–400 times higher at low N than with adequate N supplies (Table 1). It did not proceed from a dilution effect, as fructose and glucose concentrations increased by 84% and 65%, respectively, at 50 μM NO3 although growth was only reduced by 30%. Increased hexose concentration in N-deficient tobacco and tomato plants has already been reported (Paul and Driscoll, 1997; Urbanczyk-Wochniak and Fernie, 2005), and this appears coherent with an overall slowing down of plant metabolism (including respiration) following N shortage. The response of organic acid concentration to N supply diverged from that of hexoses. In this trial, acid concentration was under the detection limit (<0.03% DW) in the lowest N treatments but increased sharply with N supply up to ∼0.6 mol NO3 m−3 in solution (Fig. 5a, Table 1). Thereafter, the concentrations continued to rise but the slope was 50 times slower. This pattern mimicked that described for leaf [NO3] (Fig. 3). Organic acid concentrations, and in particular that of malic acid, are known to evolve in response to the cation–anion plant uptake imbalances, in particular those provoked by altered NO3 supplies, and to the control of cytoplasmic pH. Textbooks (such as Mengel and Kirkby, 1987) described how the pH-stat model of Davies enables malate synthesis to counter the internal pH rise associated with NO3– assimilation. This behaviour was described for tomato plants fed with increasing solution [NO3] (Kirkby and Knight, 1977). Furthermore, nitrate acts as a signal to initiate a coordinated increase in the expression of different genes involved in organic acid synthesis, leading to accumulation in N-sufficient tobacco plants (Scheible et al., 1997). Recently, leaf organic acid profile analysis of N-deficient tomato plants confirmed this behaviour for the major acids of the tricarboxylic acid (TCA) cycle such as malate, citrate, iso-citrate, fumarate, succinate, and 2-oxoglutarate (Urbanczyk-Wochniak and Fernie, 2005). In the present trial, it is noteworthy that the amount of organic acid C reserves is similar to that of hexoses, but their responses to N limitation are opposite. Therefore, it may be remarked that carboxylic acid accumulation with rising N supply nearly compensates for depletion of the hexose store. This is important in growth models because the C reserves are solely ascribed to the pool of non-structural sugars, although they should also rely on other C sources. Leaf phenolic concentrations responded strongly to N nutrition, as hexoses did. In the N-limiting growth domain (i.e. <300 μM), phenolic concentration increased with nitrate limitation, up to 2-fold for chlorogenic acid and rutin in the 50 μM NO3 treatment compared with adequate nutrition (Fig. 5b). There is a general agreement in the literature that N deficiency stimulates phenolic synthesis in tomato leaves (Wilkens et al., 1996b; Stout et al., 1998; Hoffland et al., 2000; Stewart et al., 2001) and in several crops (Haukioja et al., 1998; Lux-Endrich et al., 2000; Kováčik and Bačkor, 2007), although the shape of this relationship is unknown. Based on this ubiquitous relationship, Cartelat et al. (2005) proposed a quick test to diagnose wheat N status from the rapid optical measurement of their leaf content of polyphenolics. Hence, the authors found a highly significant negative correlation between leaf polyphenolic and N contents, independent of the growth stage. This allowed the rapid estimation of the crop nitrogen nutrition index (NNI; see Lemaire et al., 2008). Indeed they observed that when NNI was ≥1 (i.e. N status was adequate for growth), leaf polyphenolics remained constant, but when NNI was <1 (N-limiting growth) they increased as leaf N decreased. This pattern is similar to what is reported on the tomato plants here. How do results and simulations compare? The proposed model adopts the very same principles used by Herms and Mattson (1992), i.e. primary and secondary metabolites compete for the C resources produced by photosynthesis, but the sink strength of the pool of primary metabolites (i.e. the maximum rate in Equation 10) depends on N availability. As the response of secondary metabolism to N and C resources is not known, it is inferred from the respective responses of photosynthesis and growth to N and C. Inevitably, the pattern of the former depends on the patterns of the latter. Herms and Mattson (1992) imposed arbitrary sigmoid responses of NAR and RGR to soil resource availability (i.e. ns in this paper). Favouring instead a rational approach, here photosynthesis and growth were related to plant [N] on the basis of existing models with experimental backing, the link with N availability in the soil solution also being made from a published model. It is clear that output simulations (Fig. 6a) differ greatly from Fig. 1. Indeed, simulated photosynthetic rates and RGR are saturable, but not sigmoid, and this is clearly driven by the N uptake submodel. Nonetheless, simulations conform to the predictions of Herms and Mattson that ‘at low to moderate level of resource availability … rates of net assimilation, growth and secondary metabolism are positively correlated’. In contrast, at a moderate to high level of resource availability, the simulations (Fig. 6a) contradict previous assumptions (Fig. 1), that ‘… relative growth rate and secondary metabolism are inversely correlated …’, although in this region photosynthesis and growth show similar patterns in Figs 1 and 6a. This discrepancy is easily explained. Indeed, C-based secondary metabolites do not contain N, but the metabolic machinery necessary for their synthesis does need and does contain N. Thus, as shown by Fig. 6b, any [N] increase benefits both primary and secondary metabolism. Unfortunately, there is no way to measure the overall rate of secondary metabolism and thus to assess the likelihood of Fig. 6a versus Fig. 1. Instead, it may be tempting to infer secondary metabolism from the measurement of metabolite concentrations. For instance, Stamp (2004) interpreted the bell-shaped curve of secondary metabolism in Fig. 1 as reflecting the concentration of secondary metabolites expressed as a percentage of plant dry biomass (Fig. 2 in the cited paper). The present experimental results are restricted to the domain of agricultural practices, i.e. zones 2 and 3 in Fig. 1. In this range, a decreasing concentration of phenolics with increasing ns up to 0.3 mol NO3 m−3 was observed, followed by a constant concentration thereafter. Such an observation conforms to Stamp's prediction and other experimental results (Stout et al., 1998; Schmelz et al., 2003; Glynn et al., 2007). However, this does not validate the initial hypothesis, because the present model predicts a decrease of [C2] with increasing ns (Fig. 6c), although the rate of secondary metabolism increases (Fig. 6a, b). Indeed, according to Table 2, the primary metabolism has much higher affinity for C resources than the secondary metabolism (i.e. K1 << K2). Thus, Wm accumulates more C than W2 when [N] increases (Fig. 6b). It results that [C2] declines only because the accumulation of primary metabolites dilutes secondary metabolites (Fig. 6d). Herms and Mattson (1992) emphasized the idea that C-based secondary compound synthesis is favoured when C resources increase in the plant stores. Thus, it seems logical to expect [C2] to be positively related to [Cc]. Figure 7 is an outcome of the model, which predicts a more complex relationship, because the correlation gets inversed from low to high [N]. It was not possible to find in the literature the data required to test this prediction. However, the present data address a mere question about the nature of these C resources. Figure 8a shows a positive relationship between tomato leaf phenolic and non-structural carbohydrate concentrations, supporting the model prediction pattern (Fig. 7) in the ‘agronomic’ domain. However, non-structural carboxylic acids are C substrates for various biosynthetic pathways and they might also be accounted for in the mean store of C resources. This would be relevant considering their strong response to N supply (Fig. 5a). Leaf phenolic concentration was also plotted against the sum of the concentrations of non-structural carbohydrates and carboxylic acids (Fig. 8b). The course of the relationship was reversed, which poses question about the nature of the C resources that should be integrated in this C pool. Fig. 8. Open in new tabDownload slide Measured leaf phenolic concentration (g per 1000 g DW) as a function of measured C resource concentration (g per 100 g DW) taken as (A) non-structural carbohydrates or (B) the sum of non-structural carbohydrates and carboxylic acids. Lines are quadratic smoothing. Fig. 8. Open in new tabDownload slide Measured leaf phenolic concentration (g per 1000 g DW) as a function of measured C resource concentration (g per 100 g DW) taken as (A) non-structural carbohydrates or (B) the sum of non-structural carbohydrates and carboxylic acids. Lines are quadratic smoothing. Conclusion The experiment confirms literature data about accumulation of primary and secondary compounds in relation to wide-ranging N nutrition. The C allocation to both pools is predictable from a simple model merely based on trophic hypotheses. It suggests that secondary compound concentration declines at high N availability following dilution by primary metabolites and not necessarily a lesser rate of secondary metabolism. Thus, it follows that plants of low N status are likely to produce a high concentration of ‘defensive’ compounds, giving them an advantage against aggressors. Conversely, this model predicts that high rates of N fertilization are likely to produce larger amounts of secondary compounds, which may represent a better strategy for production purposes. It is a challenging issue to infer the likelihood of this prediction, but more experimental work is obviously required. Abbreviations Abbreviations A total leaf area [C2] concentration of C-based secondary compounds [Cc] concentration of non-structural carbon resources [Cm] concentration of metabolic compounds das days after sowing DW leaf dry biomass GAR gross CO2 assimilation rate GDB growth–differentiation balance GDBH growth–differentiation balance hypothesis [N] total nitrogen concentration NAR net CO2 assimilation rate NFT nutrient film technique NNI nitrogen nutrition index [NO3] nitrate concentration PPFD photosynthetic photon flux density RGR plant relative growth rate We thank J Fabre, J Hostalery, and V Serra for conducting the experiments, D Vailhen and P Orlando for help with harvests, and E Rubio and D Bancel for chemical analyses. We are grateful to Dr DJ Pilbeam (University of Leeds, UK) for English revision and valuable comments. References Adamowicz S , Le Bot J . 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