Blackwell Science, LtdOxford, UKPCEPlant, Cell and Environment0016-8025Blackwell Science Ltd 2005? 2005 28?14211435 Original Article Plant primary metabolism during defence J. Scharte et al.
Plant, Cell and Environment (2005) 28, 1421–1435
Photosynthesis and carbohydrate metabolism in tobacco leaves during an incompatible interaction with Phytophthora nicotianae JUDITH SCHARTE*, HARDY SCHÖN* & ENGELBERT WEIS
Institut für Botanik, Westfälische Wilhelms-Universität, Schlossgarten 3, 48149 Münster, Germany
ABSTRACT The metabolic and cellular changes in source leaves of Nicotiana tabacum L. cv SNN during an incompatible interaction with Phytophthora nicotianae van Breda de Haan were investigated and compared with defence reactions. Hypersensitive cell death was preceded by a rapid and highly localized shift to non-assimilatoric metabolism. During the first 6 h post infection (hpi), reactive oxygen species (ROS) accumulated. Callose was deposited at the interface of adjacent mesophyll cells (≥1 hpi), the export of sucrose collapsed and its content in the apoplast increased. Stomata closed and photosynthetic flux was reallocated from CO2 assimilation in favour of photorespiration. This was accompanied by an increase in respiration, glucose-6-phosphate dehydrogenase (G6PDH) activity, apoplastic invertase and hexose content. Later (>6 hpi) the photosynthetic electron transport chain was interrupted and photosynthesis completely collapsed. This was accompanied by a further increase in apoplastic invertase and carbohydrates, respiration and oxidative pentose phosphate pathway (OPPP) and followed by further burst in ROS release. Hypersensitive cell death did not appear until photosynthesis completely declined. Photosynthesis was visualized by chlorophyll-a fluorescence imaging on a macro- and microscopic scale. Decline in photosynthesis and defence reactions were highly localized processes, which occur in single mesophyll cells. We propose that in photoautotrophic leaves, photosynthesis and assimilatory metabolism must be switched off to initiate respiration and other processes required for defence. An early blockage of intercellular sugar transportation, due to callose deposition, in conjunction with enhanced apoplastic invertase activity could facilitate this metabolic shift. Key-words: Nicotiana; chlorophyll-a fluorescence imaging; hypersensitive response; oxidative pentose phosphate
Correspondence: Judith Scharte. Fax: +49 2518323823; e-mail:
[email protected] *These authors contributed equally to this paper. © 2005 Blackwell Publishing Ltd
pathway (OPPP); plant defence; primary metabolism; reactive oxygen species.
INTRODUCTION In plant–pathogen interactions, disease resistance is generated by rapid activation of a multitude of defence reactions, including reinforcement of the cell wall (for review, see Lamb & Dixon 1997), phytoalexin production (for review, see Hammerschmidt 1999), expression of various pathogenesis-related (PR) proteins (Van Loon & Van Strien 1999 and references therein) and the generation of ROS, such as superoxide anion radicals and hydrogen peroxide (Apel & Hirt 2004; and references therein). Usually, the hypersensitive defence reaction leads to cell death. These processes are associated with increased demands for energy, reducing equivalents and carbon skeletons for biosyntheses. Whereas respiration, OPPP and the shikimic acid pathway are usually enhanced in infected tissue (e.g. Scheideler et al. 2002), photosynthesis can initially be stimulated, but is often depressed at later stages of an infection (e.g. Bassanezi et al. 2002). A close linkage between plant carbohydrate-status and the outcome of a plant pathogen interaction is evident. It includes the phenomenon of ‘high sugar resistance’ (Horsfall & Dimond 1957) and the finding that various pathogenesis-related genes are sugar inducible (e.g. Herbers et al. 1996a, b). Overexpression of a yeast invertase in the plant apoplast results in expression of PR proteins and increased resistance against virus infection (Herbers et al. 1996a, b, 2000). The well-documented induction of apoplastic invertase activity by various biotic stresses (Roitsch et al. 2003; and references therein) actually suggests a co-ordinated regulation of plant source/sink relations and the response to pathogens (Ehness et al. 1997). Carbohydrate retention is favourable for plant defence, but may also support the parasite (e.g. Ayres, Press & Spencer-Phillips 1996). Thereby, sugars play a dual role for a successful plant defence, as fuel for metabolic pathways and as regulators of gene activity via sugar sensing. Soluble hexoses are known to repress source-specific genes, such as those for photosynthesis, and activate sink-specific genes, as those for respiration, OPPP and shikimic acid pathway (for review, 1421
1422 J. Scharte et al. see Rolland, Moore & Sheen 2002). Nevertheless, the contribution of primary metabolism to the plant disease resistance is still poorly understood (Conrath et al. 2003) and a consistent concept of the inter-relationship between changes in primary metabolism and a successful plant defence does not yet exist. The metabolic situation in the photoautotrophic mesophyll is not well suited for successful defence. Secondary- and dark-metabolism are tuned down, while photosynthates are rapidly exported. Some metabolic processes, such as respiration and OPPP, required for defence, may even be in conflict with the metabolic requirements for photosynthetic assimilation. For example, Calvin cycle and OPPP in the stroma of the chloroplasts are complementary regulated via the thioredoxin system to avoid futile cycling (Scheibe 1991). Here, we examined a multiplicity of alterations in plant primary metabolism in source leaves of tobacco – such as sugar export, carbohydrate status, stomatal aperture and photosynthesis – during an incompatible interaction with Phytophthora nicotianae (P. nicotianae). This oomycete is a soil-borne, hemibiotrophic plant pathogen with a broad host-range of over 70 tropical and temperate crops, predominately solanaceous plants (Erwin & Ribeiro 1996 and references therein). It causes the Black Shank disease of tobacco, which includes root rot, leaf wilting, stem blackening and eventual death. Unique features of oomycetes, such as none or few membrane sterols (Judelson 1997), which are targets for toxic saponins during plant defence against filamentous fungi (Osbourn 1996), suggest that plants must possess distinct genetic and biochemical mechanisms to fend off oomycetes. Until now there is little information about defence-related changes in host primary metabolism during an infection with oomycetes. We present evidence for early metabolic and cellular transitions at the infection site, eventually leading to a complete decline in photosynthesis. These changes precede, and are possibly an important prerequisite for, defence and hypersensitive cell death. We could visualize most of these alterations at a macroscopic and microscopic scale using chlorophyll-a fluorescence imaging and demonstrate, that these processes remain highly localized and can occur in single mesophyll cells.
MATERIALS AND METHODS
Zoospores production and inoculation Zoospores were produced under aseptic conditions according to von Broembsen & Deacon (1996). Finally the zoospores were stored in sterile tap water. Small areas of source leaves were infiltrated with a suspension containing 500–1000 zoospores mL-1 (‘zoospore-leaf infiltration assay’, Colas et al. 2001). Control tissues were infiltrated with sterile tap water. Inoculation of the plants was always performed 1 h after beginning of the photoperiod.
Imaging of photosynthetic electron transport from chlorophyll-a fluorescence Mapping of photosynthetic electron transport (PET) was derived from the chlorophyll-a fluorescence imaging system, connected with a gas exchange system as described in Siebke & Weis (1995b) and Jensen & Siebke (1997). Attached leaves were placed in a sandwich-type gas exchange cuvette. During measurements the leaf temperature was maintained at 22–24 ∞C. CO2- and O2-concentration in the gas phase were adjusted as indicated in the text. The internal CO2 could be calculated from gas exchange, according to Siebke & Weis (1995b). Images of PET (mmol electrons m-2 s-1) were calculated pixel-by-pixel from fluorescence images, basically as described in Jensen & Siebke (1997) and Meng et al. (2001). According to Schreiber (1997) we assumed 84% mean light absorption by the leaf. Photosynthetic induction was measured during the first 2 min of illumination (280 mmol quanta m-2 s-1) after 1 h of dark adaptation. From PET images, taken during the first 120 s of illumination, an ‘integral induction image’ was calculated according to Meng et al. (2001), which represents the sum of electrons transported during that period.
Gas exchange Attached leaves were positioned in a gas exchange cuvette as described above. Gas exchange was measured with a two-channel gas flow system, as described in Siebke & Weis (1995a). The gas flow rate through the chamber was 1000 mL min-1; the leaf temperature was maintained at 22– 24 ∞C and the relative humidity at 55–57%. CO2- and O2concentrations were adjusted using gas flow controllers (Tylan General GmbH, Eiching, Germany).
High-resolution imaging
Plant and oomycete material Tobacco plants (Nicotiana tabacum L. cv SNN) were grown in a growth chamber with 24 ∞C/22 ∞C day/night temperature and 14 h photoperiod (300 mmol quanta m-2 s-1). Sixto 8-week-old plants at the five-leaf stage were used for the analysis. Phytophthora nicotianae van Breda de Haan isolate 1828 (DSMZ GmbH, Braunschweig, Germany) was cultivated at 24 ∞C on clarified V8-Agar as described by von Broembsen & Deacon (1996).
Chlorophyll-a-fluorescence imaging of single cells was accomplished as basically described by Oxborough & Baker (1997). Basis for our imaging system is a Leica DMRBE microscope (Leica, Wetzlar, Germany). For highresolution imaging we used Leica PL FLUOTAR lenses 10¥/0.30 and 25¥/0.75 OIL. A Leica 105z 50 W mercuryvapour lamp provided irradiance for actinic and saturating light. Intensity of radiation was adjusted by a set of neutraldensity filters (Schott NG; Schott AG, Mainz, Germany). Excitation light was filtered by an Schott BG 39 and
© 2005 Blackwell Publishing Ltd, Plant, Cell and Environment, 28, 1421–1435
Plant primary metabolism during defence 1423 deflected by a dichroic mirror (585DCRX; Chroma, Irvine, CA, USA), while emitted chlorophyll-a-fluorescence passed an Schott RG645 LP and was recorded by a highperformance charge-coupled device (CCD)-camera (12 bit SensiCam; PCO AG, Kelheim, Germany). Photosynthetic active irradiance was determined according to Küpper et al. (2000). PET images were calculated pixel by pixel, basically as described by Siebke & Weis (1995a, b) using Optimas (Media Cybernetics, Silver Spring, MD, USA).
P700 redox kinetics P700 redox kinetics were recorded with a PAM 101 spectroscope with the emitter-detector-unit ED-P700-DW (810– 860 nm) (Heinz Walz GmbH, Effeltrich, Germany) according to Klughammer & Schreiber (1998). Measurements were performed in a reflecting leaf chamber with leaf discs, infiltrated with a 10-mM methylviologen (Paraquat) solution and preincubated for 5 min. Signals were measured repetitively with 20 s intervals.
Total RNA extraction and northern-blot hybridization Total RNA was extracted according to Logemann, Schell & Willmitzer (1987). RNA (7 mg) was subjected to electrophoresis in a 1% agarose-formaldehyde gel, capillary transferred and linked onto Hybond N+ membrane (Amersham Biosciences Europe GmbH, Freiburg, Germany). Probes were labelled with biotin by polymerase chain reaction according to the method described by Löw & Rausch (1996). Hybridization was performed in a reaction mixture containing 50% (v/v) formamide, 1% (w/v) sodium dodecyl sulphate, 1 M NaCl, 6% (w/v) PEG 6000 and 250 mg mL-1 salmon sperm DNA at 42 ∞C. Membranes were washed with 0.2 SSC/0.5% (w/v) SDS at 65 ∞C, developed as described by Löw & Rausch (1996) and exposed to Hyperfilm ECL (Amersham Biosciences Europe GmbH, Freiburg, Germany).
Histochemical detection of H2O2 and Callose Hydrogen peroxide (H2O2) accumulation was detected by in situ staining with 3,3-diaminobenzidine (DAB) following a modified protocol from Thordal-Christensen et al. (1997) Leaves were placed for 6 h in DAB solution (1 mg mL-1, pH 3.8). The incubation was stopped in boiling ethanol (96%, v/v) for 10 min. A dark-brown polymerization product indicates the DAB reaction with H2O2. Callose was stained by the aniline blue method as described by GómezGómez, Felix & Boller (1999). Stained sections were viewed using a fluorescence microscope (Leica DMRB) equipped with a filter set (340–380 nm excitation filter, 400 nm dichroic mirror, 425 nm barrier filter).
Leaf epidermal peels Epidermal peels were taken from the adaxial side of a leaf using fine forceps (Inox no. 4; Dumont and Fils, Montignez,
Switzerland) and mounted on a coverslide in distilled water. The stomatal aperture was determined within 10 min after preparing the peels with a microscope (Leica DMRB, Leica, Wetzlar, Germany).
Chlorophyll determination Leaf discs from inoculated or control areas were rapidly frozen in liquid nitrogen, ground to a powder and the chlorophyll was extracted with 80% (v/v) acetone. Total chlorophyll content was determined as described by Porra, Thompson & Kriedermann (1989).
Extraction of apoplastic fluid Preparation of apoplastic fluid was carried out as described by Lohaus et al. (2001) with the following modifications: Plant tissues were infiltrated with 1 M NaCl, the surface was dried and the leaf discs were centrifuged at 2500 g for 10 min at 4 ∞C in a 25-mL syringe barrel placed in a centrifuge tube. The apoplastic fluid was frozen in liquid N2 and stored at -70 ∞C until measurement.
Photometric determination of G6PDH- and invertase activity, carbohydrate content and sucrose efflux Freshly cut leave discs were frozen in liquid N2 and ground to a fine powder. Extraction and determination of G6PDH activity was accomplished as described by Fickenscher & Scheibe (1986). The invertase activity in the apoplastic fluid was determined as described by Jones, Outlaw & Lowry (1977) and the content of apoplastic hexoses according to Kunst, Draeger & Ziegenhorn (1984). For sucrose efflux measurements through the petiole whole leaves were fully inoculated with either zoospores or water. The measurements of sucrose efflux were performed according to Murillo et al. (2003).
RESULTS Hypersensitive response We infected source leave of N. tabacum with P. nicotianae by infiltrating small mesophyll areas with a suspension of zoospores (‘infection site’) (Fig. 1a). We have chosen this zoospore-leaf infiltration assay (Colas et al. 2001) to achieve a rapid, synchronized start of infection of all parenchymatic cells in the infiltrated area. For mock-inoculation we infiltrated sterile tap water instead of zoospores, further named as ‘control’. To consider individual variations between different plants and leaves, the samples for control and infection site were taken from adjacent intercostal areas of the same source leaf. During the first hours post infection (0–12 hpi), prior to the formation of macroscopic visible necrosis, ROS (= 1 hpi, Fig. 1b) and transcripts of the pathogenesis-related protein PR-Q (Fig. 1c) locally accumulate at the infection site. PR-Q, encoding for an acid chitinase (Payne et al. 1990), has been found to be induced
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Figure 1. Defence reactions of Nicotiana tabacum L. cv SNN infected with Phytophthora nicotianae. (a) Images of an infiltrated tobacco leaf. At 0 hpi: Leaf areas infiltrated with water (control) or zoospores (infection site). At 48 hpi: Hypersensitive lesions appear at the infection site. (b) DAB staining, indicating H2O2 release at the infection site (1 hpi). (a) and (b) White bar: 10 mm. (c) Gene expression of the pathogenesis-related protein PR-Q. Total RNA was isolated from control and infection site areas of a source leaf (12 hpi). Probing with an 18S cDNA served as loading control.
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Figure 2. Defence-induced formation of callose and H2O2 accumulation. Leaves were stained with aniline blue and examined under UV light. Bright green fluorescence indicates callose deposition. (a) Infection site, 1 hpi; (b) infection site, 6 hpi and (c) control, 3 hpi. (d to f) Localization of callose formation in mesophyll tissue at the infection site: (d) 1 hpi, (e) 3 hpi, and (f) 6 hpi. (g to i) Callose deposition (green fluorescence) and H2O2 accumulation (DAB, brown staining) in spongy mesophyll cells at 3 hpi. (g and h) infection site and (i) control. White bars: (a to c) 500 mm, (d to i) 100 mm. (a), (d) and (g): White arrows point to callose deposition at cell-to-cell contacts of infected cells. Black arrows indicate cell-to-cell contacts in control tissue, ic the intercellular space. Images (c) and (i) are representative for all controls. © 2005 Blackwell Publishing Ltd, Plant, Cell and Environment, 28, 1421–1435
Plant primary metabolism during defence 1425 by salicylate (Ward et al. 1991) and may therefore be taken as an indicator for systemic defence response. Neither PR-Q transcripts nor ROS release could be detected in the control (Fig. 1b & c). This indicates that there was no significant systemic response during this phase of defence. From 12 hpi onward, the infected tissue became chlorotic and collapsed. Eventually, a sharply bounded hypersensitive lesion was formed, exactly congruent to the initially infiltrated leaf area (Fig. 1a). Over a period of 4 weeks after infection the necrotic lesions did not spread beyond the initial inoculation site. Neither did any indications of the development of an oomycete mycelium nor symptoms of the Black Shank disease became visible. Therefore, we regard the interaction as incompatible. In the following, we describe local metabolic and defence induced alterations at the infection site during the first 12 hpi, preceding chlorosis and cell death.
Defence-induced changes in carbohydrate metabolism Callose formation Already at 1 hpi, callose formation could be observed at the infection site, which proceeded over the course of infection (Fig. 2a & b), but never occurred in the control (Fig. 2c). Initially, callose distinctively appeared at cell-tocell contacts of spongy mesophyll cells (Fig. 2d & e). During the following hours callose spread over the whole cell wall of infected cells (Fig. 2f). Using combined aniline blue-/DAB-staining we found, that almost all cells at the infection site producing H2O2 also exhibit callose deposition at the interfaces to adjacent cells (Fig. 2g & h). DAB staining never significantly spread across the callose barriers. The ROS accumulation also appears to be localized in single mesophyll cells (Fig. 2g & h), but does never appear in the control (Fig. 2i).
Decline in sucrose efflux During the first hours post infection, the export of sucrose from the infected mesophyll through the petiole decreased about 70% compared to control leaves (Fig. 3a). Simultaneously, the content of apoplastic sucrose increased by more than 50% in comparison to the control. During the following hours of infection (>6 hpi) the sucrose content further increased about 250% of the control (Fig. 3b).
Apoplastic invertase and hexoses The increase in the content of apoplastic sucrose was accompanied by enhanced expression (Fig. 3c) and activity (Fig. 3d) of the apoplastic invertase at the infection site. In control tissue, transcripts of apoplastic invertase were not detectable and the invertase activity was low. At the infection site, the level of the apoplastic invertase transcripts continuously increased until 6 hpi (Fig. 3c), while the activity of the enzyme rapidly increased with a first maximum
at 2 hpi. From 4 hpi on, a second increase by more than 250% of the control was found (Fig. 3d). A comparable increase in invertase activity has been detected after elicitation with a cell-free filtrate of P. nicotianae zoospores, suggesting, that the major part of the activity can be attributed to the plant (Fig. 3d). During the first gain of invertase activity (1–4 hpi), the content of apoplastic hexoses increased only moderately, but increased parallel to the invertase activity at later stages of infection (Fig. 3e). Elicitation with a cell-free filtrate of P. nicotianae zoospores has the same effect in the content of apoplastic hexoses, as the infection with zoospores (data not shown).
Stimulation of respiration and oxidative pentose phosphate pathway Respiration, measured as CO2 production in the dark, was strongly stimulated (about 70%) during the first hour post infection (Fig. 4a). After stagnation until 9 hpi a second increase of about 250% compared to the control was detected. The stimulation of respiration may be even underestimated, as small parts of uninfected tissue could be included in the gas exchange cuvette. Leaf tissue, elicited by cell-free filtrate of the pathogen, exhibited a comparable stimulation in dark respiration (data not shown), suggesting that enhanced CO2-uptake mainly reflects an increase in plant respiration. G6PDH, a key enzyme of the OPPP, also exhibited a rapid initial increase of activity (1 hpi, about 40%), and a further one with a maximum between 6 and 10 hpi (Fig. 4b). In plants, the OPPP is known to be located in the cytosol as well as in the stroma of the chloroplasts. Metabolites can be exchanged through triose-phosphate carriers in the chloroplast envelope and equilibrate between cytosol and stroma (for review, see Flügge 1999). In the stroma, the OPPP shares metabolites with the Calvin cycle (Schnarrenberger, Flechner & Martin 1995). Thus, any enhancement of the OPPP in the dark and related increases in the stromal level of metabolites of the OPPP are expected to stimulate the initiation of the Calvin cycle and, hence, the induction of photosynthesis after a dark period. Fast photosynthetic induction zones have actually been observed in developing tissue at the base of expanding tobacco leaves, where they mark carbohydrate consuming sink-zones and high stromal levels of OPPP intermediates (Meng et al. 2001; and references herein). We analysed the photosynthetic induction during the first 2 min after a 1-h dark period. Measurements of photosynthetic induction were performed in 21% O2 and 65 mL CO2 L-1 (close to the CO2-compensation point). Under such experimental conditions, stomatal aperture was proved to have no effect on the photosynthetic induction (data not shown). Using chlorophyll-a fluorescence imaging, we could visualize the photosynthetic induction (Fig. 4c) and found, that photosynthetic induction was accelerated in infected leaf patches (where the G6PDH activity was high, Fig. 4b), but remained slow in the surrounding tissue. Zones of fast pho-
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Figure 3. Time courses of defence-induced changes in carbohydrate metabolism. (a) Repression of sucrose efflux at the infection site. The insert shows the total amount of exported sucrose (open circles: control; closed circles: infection site). (b) Increase of apoplastic sucrose content at the infection site. The mean value of apoplastic sucrose content in controls was about 135 mmol m-2. (c) Gene expression of apoplastic invertase. (d) Enhanced apoplastic invertase activity after infection and elicitation with a cell-free filtrate of Phytophthora nicotianae zoospores. (e) Increase of apoplastic hexose content at the infection site. The mean value of apoplastic invertase activity in the controls was about 394 nKat m-2 and that of the hexose content about 63 mmol m-2. All percentage increases are values of relative activities or contents based on the control (= 0%). Values are means ± SE (n = 4).
tosynthetic induction were strictly bounded and exactly congruent to the initial infection zones.
Stomatal closure and inhibition of photosynthesis We examined the stomatal aperture by microscopic evaluation of freshly prepared epidermal peels. The fraction of open stomata (all stomata ≥3 mM) strongly decreased at the infection site during the first 6 hpi, but increased again at later stages of the infection (Fig. 5a), while stomata in the controls remained open over the whole investigation period. From gas exchange measurements, we calculated a decline in intercellular [CO2] (Ci) from about 250 in uninfected to about 120 mL CO2 L-1 in infected tissue. At 6 hpi, the stomatal conductance for water (gH2O) decreased in infected leaf areas by about 42% compared to control leaves: control, 111.2 ± 23.0 (mmol m-2 s-1); infection site (6 hpi), 64.4 ± 14.8; values are mean ± SE (n = 4). This decline in gH2O could even be underestimated, as we could not exclude some contribution of uninfected parts of the mesophyll to the gas exchange measurements. As a consequence of stomatal closure, the rate of CO2 dependent PET decreased. This was analysed by measuring the photosynthetic electron transport from chlorophyll-afluorescence images, taken in the presence of low (2%) O2,
enhanced CO2 concentration (700 mL CO2 L-1) and nearly saturating light (820 mmol quanta m-2 s-1; Fig. 5B & C). Under such conditions, photorespiration is largely suppressed (while mitochondrial respiration is not) and electrons from PET are largely consumed by CO2 fixation (Sharkey, 1988). In control areas, PET rates in 2% O2 of about 110 mmol electrons m-2 s-1 were calculated. At the infection site, the photosynthetic flux in 2% O2 rapidly collapsed (Fig. 5B & C). The flux reached a preliminary plateau between 3 and 6 hpi before it further declined. In the presence of 21% O2, when electrons are consumed either by CO2 fixation or photorespiration, only a weak inhibition was been during the first 6 hpi (Fig. 5B & D), but later the photosynthetic flux equally decreased in the presence or absence of high O2 (Fig. 5B–D). The O2 sensitive inhibition of PET (1 to 6 hpi) indicates the inhibition of photosynthetic CO2 fixation, as a result of stomatal closure. The O2 insensitive decline occurred when the stomata opened again (>6 hpi) and reflects a general decrease in the photosynthetic capacity of the infected mesophyll. Again, the inhibited patches were congruent to the initial infection zones. The stoma-dependent decline in photosynthesis (in 2%O2) occurred at a somewhat broader and gradually bounded area, compared to a more strictly bounded inhibition area seen in 21% O2 (compare Fig. 5C to Fig. 5D). This is consistent with the suggestion, that in 2% O2, the
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Integrated PET (µmol electrons m–2 2 min–1)
Figure 4. Defence-induced changes in dark-metabolism. Increase of (a) dark respiration and (b) glucose-6-phosphate dehydrogenase activity at the infection site, expressed as percentage increase relative to values from the control area in the same leaf. The inserts show the original data from control (open circles) and infected (closed circles) areas. (a and b) All percentage increases are values of relative activities based on the control (= 0%). Values are means ± SE (n = 4). (c) Images of the relative rate of photosynthetic induction (integrated photosynthetic flux during the first 2 min of photosynthetic induction). Low values (blue) indicate relatively slow photosynthetic induction; high values (green – red) indicate stimulation in photosynthetic induction. White bar: 10 mm.
decline in flux is due to reduced stomatal conductance. Even if stomatal closure is strictly localized, the resulting low CO2zone would be somewhat more diffuse, because of lateral gas exchange within the intercellular space of the mesophyll.
Inhibition of photosynthetic electron transport At an infection stage, when stoma-independent photosynthesis was already severely reduced, the mesophyll at the infected sites was fully turgescent and still did not show any visible symptoms of the hypersensitive reaction (HR). The chlorophyll content (Fig. 6a) and chlorophyll a/b ratio (data not shown) were not or only little affected.
Significant loss in chlorophyll – mainly chlorophyll a – appeared at a later stage (>12 hpi; data not shown). This is in agreement with the observation that the transcripts of chlorophyll a/b binding proteins (cab) at the infection site remained more stable, compared to the Rubisco smallsubunit (rbcS), and did not disappear before 12 hpi (Fig. 6b). To test the activity of the electron transport chain in situ, we infiltrated the leaves with the electron acceptor methylviologen and followed the reduction-kinetics of P700+, the oxidized chlorophyll-dimer in the reaction centre of PSI, after a saturating light pulse. In the presence of methylviologen, P700 was fully photo-oxidized during a 120-ms saturating light pulse. At 8 hpi, the amplitude of the P700 signal,
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Figure 5. Defence-induced changes in stomatal aperture and photosynthesis. (a) Time course of stomatal closure at the infection site (percentage of open stomata, relative to total stomata in the inspected leaf area) open circles: control; closed circles: infection site. Values are means ± SE (n = 3). For every point in time and every sample a total number of 90 ± 20 stomata was counted. (b) Changes in capacity of PET due to changes in gas-composition from 700 mL CO2 L-1, 21% O2 (total photosynthetic flux – circles) to 700 mL CO2 L-1, 2% O2 (assimilatory flux – triangles). (c and d) PET-images (steady-state flux) taken at different times after infection. Images were taken in (c) 2% O2 or (d) 21% O2. White bar: 10 mm.
indicating the relative number of photoactive PSI centres, was not or only slightly affected (Fig. 6c, inset). However, the dark relaxation of the signal, indicating P700+ reduction kinetics, was slowed down, indicating that the electron transport chain was restricted upstream of PSI. The rate-constant of electron donation to P700 decreased from 0.041 (ms-1) in healthy mesophyll to values below 0.025 (ms-1) at 8 hpi (Fig. 6c). This suggests that an inhibition of the photosynthetic electron transport chain contributes to the overall decline in photosynthesis. We cannot exclude that reactions of the photosynthetic carbon metabolism, such as the Rubisco activity, are also slowed down and contribute to the overall decline in photosynthesis. Actually, we do see a rapid decline in rbcS transcripts at the infection site (Fig. 6b). However, we did not yet test whether the decline in transcripts actually leads to a decline in the Rubisco activity.
Correlation of local changes in plant metabolism and HR Figure 7a shows the initial infiltration with the pathogen (infection site; upper part of the leaf), and tap water (control; lower part of the leaf). Figure 7b–e exhibits the spatial
distribution of local stimulation of the OPPP – indicated by enhanced photosynthetic induction – (Fig. 7b, 3 hpi), local stoma-dependent (Fig. 7c, 3 hpi) and -independent decline in photosynthesis (Fig. 7d, 6 hpi), and H2O2 accumulation (Fig. 7e, 9 hpi). Evidently, the initial infection site was exactly congruent to the site of metabolic changes and ROS accumulation. The initial inoculation zone was also congruent to the final hypersensitive lesion (Fig. 1a). This indicates, that all changes described here, occurred in a strictly localized way, and no systemic propagation of these reactions were seen. The strict localization and congruency of the decline in photosynthesis and HR has also been visualized with microscopic resolution (Fig. 7f & g). Even if the metabolic changes appear to occur homogenously at the infection site, microscopic resolution allowed us to detect differential reactions of single mesophyll cells (Fig. 7f & g), comparable to callose and ROS formation in single cells (compare with Fig. 2). Figure 7f shows the photosynthetic activity of single mesophyll cells in situ at the border of an infection site (7 hpi). Photosynthetically active or partially inhibited cells were adjacent to fully inactive cells. Figure 7g shows the same section as in Fig. 7f under UV excitation after the elution of chlorophyll. Photosynthetically active
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Figure 6. Defence-induced changes in photosynthetic electron transport chain. (a) Chlorophyll (a + b) content of control (open circles) and infection site (closed circles). Values are means ± SE (n = 3). (b) Transcripts of chlorophyll a/b binding protein (Cab) and Rubisco smallsubunit (rbcS). (c) The rate of photosynthetic electron transport (PET) versus the rate-constant of P 700+ reduction (after a saturating light pulse). PET was measured in 21% O2 from infected leaf areas at different times after infection (compare to Fig. 5b). P700 kinetics was measured from these leaves in the presence of methylviologen. The inset shows two original P 700 transients – photo-oxidation by a 120-ms light pulse, followed by reduction in the dark – for control and infection site (8 hpi). The number beneath each data point denotes the time after infection (hpi).
cells at the infection site and in the control exhibited a bluish fluorescence, which may derive from phenolic acids (yellow-marked cells in Fig. 7f & g). We also observed cells, still exhibiting blue fluorescence, but with largely inhibited photosynthesis (white-marked cells in Fig. 7f & g). A smaller number of non-photosynthesizing cells exhibited bright yellow-white autofluorescence (red-marked cells in Fig. 7f & g). At a later stage of infection, this autofluorescence became dominant. It most likely reflects cell wall fortification (Tiwari, Belenghi & Levine 2002) and subsequent cell death. The bright autofluorescence was preceded by a light red-brown fluorescence. We do not know the origin of that brown fluorescence, but it most likely represents a transitory state of the beginning cell wall fortification. A comparison of the PET- and UVexcited images (Fig. 7f & g) demonstrates that autofluorescing, and non-photosynthetic cells could be located directly adjacent to photosynthetically active cells. Obviously, the described metabolic alterations are pass–fail reactions and their occurrence is strictly localized in single infected mesophyll cells.
DISCUSSION The metabolic situation of photoautotrophic source leaves is not well suited for defence. Carbohydrates produced during photosynthesis are rapidly exported, while metabolic pathways such as glycolysis, OPPP and shikimic pathway,
which are important for defence, are tuned down. Here we describe a complex metabolic scenario, which demonstrates that plant defence is preceded, maybe facilitated, by a fundamental shift of the primary metabolism towards carbohydrate consuming, non-assimilatory (heterotrophic) pathways. During the first hour of an infection, early ROS accumulation (Figs 1b, 2g & h) is accompanied by callose formation (Fig. 2a–h), collapse in sucrose export (Fig. 3a) and increase in apoplastic sucrose content (Fig. 3b). It suggests that immediately after pathogen recognition carbohydrates are retained at the infection site. Early callose synthesis is a cellular defence reaction caused by the disturbance of intracellular calcium homeostasis (Kartusch 2003). In tobacco cells, the Phytophthora elicitor cryptogein was shown to induce an early calcium influx (Tavernier et al. 1995). A number of reports have focused on the role of callose against systemic propagation of pathogenic viruses. Callose has also been found to block plasmodesmata during infection by the oomycete Phytophthora sojae (Enkerli, Hahn & Mims 1997). In our system, two observations point to callose as a general transportation barrier: (1) callose distinctly appears at the interface between spongy mesophyll cells, where intercellular metabolite exchange occurs (Fig. 2d–f); (2) H2O2, a diffusible oxygen species, spreads throughout the cytoplasm of single cells, but never spills over intercellular callose barriers, demonstrating its ‘sealing-effect’ on small diffusible molecules (Fig. 2g & h).
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1430 J. Scharte et al.
(a)
(b)
(c)
(d)
(e)
(g)
(f)
Figure 7. Congruency of local inoculation, alterations in photosynthesis, ROS accumulation and hypersensitive cell death. Spatial distribution of: (a) the initial inoculation site (infiltration zone) (0 hpi); (b) stimulated photosynthetic induction, indicating elevated OPPP intermediates (3 hpi, compare Fig. 4c); (c) decline in the rate of stoma-dependent photosynthetic CO 2 assimilation at 3 hpi (PET in 2% O2, as in Fig. 5c); (d) decline in the capacity of photosynthetic electron transport at 6 hpi (compare Fig. 5d); and (e) H 2O2 accumulation (9 hpi). (a to e) White bar: 10 mm. (f) Microscopic image of spatial distribution of photosynthetic electron transport at the periphery of an infection site (7 hpi, compare Fig. 5d). (g) UV-excited fluorescence – after elution of chlorophyll- of the same leaf-area as in image (f). Bright yellowish auto-fluorescence indicates of cell wall lignification/cell death. The brownish fluorescence most likely represents a transitory state of the beginning cell wall fortification/lignification at early stages of hypersensitive cell death. Bluish fluorescence is characteristic for uninfected, healthy cells. Yellow markings: cells with high photosynthetic rates and no signs of lignification/cell death; red markings: dead, lignificated cells (high autofluorescence) and no photosynthesis; white markings: cells with largely reduced photosynthesis and beginning of lignification/ cell death. White bar: 100 mm.
Callose deposition in the plasmodesmal neck region is actually known to retard the intercellular transport of lowmolecular weight compounds, ions and carbohydrates (Olesen & Robards 1990). We therefore conclude that defence-induced callose restricts symplastic and apoplastic export from each single elicitated mesophyll cell and, thus, export from the leaf (Fig. 3a). The increase in sucrose (Fig. 3b) was accompanied by a stimulation of the apoplastic invertase (Fig. 3d). We have made similar observations after elicitation of tobacco source leaves with NPP1 (data not shown), a necrosisinducing Phytophthora protein with structural homologues
in fungi and bacteria (Fellbrich et al. 2002). A comparable increase in apoplastic invertase activity and sucrose content was also observed after viral infection of tobacco (Herbers et al. 2000). Although low levels of apoplastic invertase transcripts already appeared shortly after infection (>0.5 hpi), the rapid initial increase in invertase activity (Fig. 3d) is likely to result from sucrose-dependent activation of the enzyme (Roitsch et al. 2003), rather than from de novo synthesis. Invertase activity, in conjunction with callose helps to retain carbohydrate at the infection site. Soluble hexoses derived from the cleavage of sucrose by the invertase reaction are re-imported into infected cells
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Plant primary metabolism during defence 1431 and fuel metabolic pathways such as respiration and OPPP. These pathways, however, are usually tuned down in photosynthesizing source leaves. This situation is turned around during defence: photosynthesis declines (Fig. 5b–d) while respiration (Fig. 4a) and the activity of G6PDH, the key enzyme of OPPP (Fig. 4b) are stimulated. A comparison of ‘photosynthesis induction images’ (Fig. 4c) and photosynthesis capacity images (Fig. 5c & d) demonstrates the antiparallel changes of photosynthesis and OPPP: the stimulation of photosynthetic induction at the infection site (Figs 4c & 7b) reflects an increased level of stromal OPPP metabolites, which feed into the Calvin cycle. At the same time, the capacity of the photosynthetic flux (Fig. 5d) declines close to zero. The decline in assimilation occurs in two steps: first by stomatal closure and later by inhibition of PET. At the infection site the stomata rapidly close (Fig. 5a) and photosynthetic CO2 uptake declines (Fig. 5b & c). Thus, in ambient O2, the overall flux remains high during the first hours (Fig. 5b & d), but is directed towards photorespiration, at the expense of carbon fixation. This allows – even in the light – metabolites of the OPPP and the glutamine–synthetase pathway to rise, while the carboxylation reaction of the Rubisco is tuned down. As a consequence, the export of triose-phosphates from chloroplasts into the cytosol decreases. Stomatal closure has often been observed during late stages of plant–microbe interactions and then seems to reflect a more general disturbance of the water status of the leaf (Nogués et al. 2002). McDonald & Cahill (1999) observed early stomatal closure, but in their system closure spread 20 mm beyond the primary infection site. Here, we observe early stomatal closure, which remains clearly restricted to the initial inoculation site (Fig. 5). As closure is preceded and accompanied by H2O2 accumulation, it could be initiated by ROS, which are known to activate calcium channels in guard cells and are involved in the osmotic responses of stomata (e.g. Pei et al. 2000). ROS as a signal for stomatal closure would also explain the localized response: callose restricts the diffusion of H2O2 (Fig. 2g & h). A significant stoma-independent depression of photosynthesis started about 4–6 hpi and rapidly proceeds within a few hours. At 9 hpi photosynthesis was close to zero, when photosynthetically inactive mesophyll was still turgescent and exhibits high rates of respiration (Figs 4a & 7f). Later, cell walls began to exhibit bright yellowish fluorescence in UV light (Fig. 7f & g). This autofluorescence has never been observed before photosynthesis was nearly depressed. Although we do not yet have direct experimental evidence, we interpret this fluorescence as an indication for lignification and hypersensitive cell death (Tiburzy & Reisener 1990). If our interpretation of these observations is correct, it indicates that photosynthesis has to be completely suppressed before the final steps of defence and cell death are initiated. It seems that mesophyll cells are regulated in such a way, that the initiation of cell wall fortification and cell death requires a ‘heterotrophic’ state of the metabolism.
Obviously, PET is a primary control step for the depression of photosynthesis. As expected for ‘high sugar cells’ (for review, see Rolland et al. 2002), rbcS transcripts disappeared at about 4 hpi (Fig. 6b). However, since the Rubisco exhibits a slow turnover, a decline of this enzyme is unlikely to cause the rapid depression of photosynthesis. Cab transcripts were more stable and did not disappear before 12 hpi (Fig. 6). Furthermore, the stability of the chlorophyll content (Fig. 6a) demonstrates that photosynthetic antennae remain relatively intact during the collapse of the photosynthetic flux. However, we do see a drastic decline in the whole chain electron transport and, in particular, in the electron transfer to oxidized PSI reaction centres, P700+ (Fig. 6c). The content of photoactive P700 (relative to chlorophyll) is little affected, suggesting that PSI itself remains stable, whereas electron donation to its centre is severely restricted. Moderate photoinhibition of PSII at the infection site (from a decline in the variable fluorescence; not shown) could be a consequence, rather than a primary cause for restricted electron transport. Recently, a drastic decline in the ‘high-potential chain’ flux (cytochrome bf Æ plastocyanin Æ PSI) was found in leaves of transgenic tobacco, exhibiting high levels of soluble sugars. This was mainly attributed to a decline in the content of plastocyanin, which seems to be an important control step in the electron transport (Schöttler, Kirchhoff & Weis 2004). Our results strongly indicate that a similar depression of the high-potential chain occurs during the stoma-independent decline in photosynthesis in plant defence (Figs 5b, d & 6c). Shortage of electron donation to PSI will prevent H2O2 release at photosystem I (Mehler reaction) and keep the stroma in an oxidized state. An oxidized state of the stromal NADPH/NADP+ and thioredoxins will keep the Calvin cycle inactive and, instead, activate stromal G6PDH, a key enzyme of the plastidic OPPP (for a review, see Scheibe 1991). An increase in G6PDH activity has actually been observed (Fig. 4b). However, the ‘induction images’ (Figs 4c & 7b) explicitly suggest an increase in stromal OPPP metabolites, which may feed into defence related biosyntheses. The shikimic acid pathway, which is well known to be activated during defence (e.g. Herrmann & Weaver 1999; Somssich & Hahlbrock 1998), is driven by PEP from glycolysis and erythrose-4-phosphate from the stromal OPPP. A sharp decline in electron transport and, consequently, in the export of triose phosphates from chloroplasts will also affect the metabolic balance in the cytosol: cytosolic fructose-2,6-bisP acts as an integrating signal for the availability of fixed carbon during photosynthesis and increases when triose phosphates decrease. We actually observed a decrease in triose phosphates in our pathosystem (data not shown). Stimulated respiratory activity will also increase fructose-2,6-bisP. Such an increase was observed, for example, after inducing wound respiration (for a review, see Stitt 1990). High levels of fructose-2,6-bisP inactivate cytosolic fructose-1,6-bisPase – involved in sucrose synthesis – and instead activate the PPi-dependent phosphofructokinase. This enzyme catalyses the cycling between triose phos-
© 2005 Blackwell Publishing Ltd, Plant, Cell and Environment, 28, 1421–1435
1432 J. Scharte et al. phates and hexose phosphates in the cytoplasm and plays a flexible role in metabolism, depending on the metabolic requirement (Hatzfeld, Dancer & Stitt 1990). Since the reaction catalysed by this enzyme is near to the equilibrium, the magnitude and direction of the cycling will respond, for example, to the needs of the cytosolic OPPP, which plays a role in the ROS production during defence, as will be discussed next. Cytosolic OPPP reactions, in particular G6PDH, were assumed to supply the plasma membrane oxidases with NADPH, by which defence-related apoplastic ROS are produced (e.g. Pugin et al. 1997). This process initiates, for example, cell wall fortification and, perhaps, hypersensitive cell death. It is not surprising, that ROS accumulation appeared quite early in our system (Fig. 1) and was maintained at a high level until cell wall lignification and cell death starts (>12 h). Maintenance of the ROS production over a period of hours requires a large amount of NADPH and carbohydrates. It is reasonable to assume that oxidation of cytosolic hexose phosphates by the G6PDH serves to supply the NADPH-oxidase with substrate (Pugin et al. 1997). Consumption of NADPH will activate the cytosolic OPPP, which, in conjunction with the fructose-2,6-Pcontrolled PPi–dependent phosphofructokinase, will maintain the flux through the NADPH generating reactions. Additionally mitochondria can function independently as a source of H2O2 (Tiwari et al. 2002), in particular during stimulated respiration, as observed in our pathosystem. Further studies must elucidate the sources for ROS and also specify the relative contribution of different ROS, such as superoxide or peroxide. As mentioned above, the accumulation of H2O2 remains strictly located to the infection site (compare Fig. 7a and e), even to single infected cells (Fig. 2g & h), and does not
spread over adjacent, uninfected mesophyll cells. Similarly, the early ‘high-OPPP state’ and the stoma-dependent and -independent decline in photosynthesis (Fig. 7) are highly localized processes, which occur at the initial infection site and remain sharply separated from uninfected cells. Photosynthetically inactive cells can even share a common cell wall with adjacent, photosynthetically active cells (Fig. 7f). Usually, the mesophyll exhibits rapid intercellular exchange of carbohydrates. Our observations point to a defence-related interruption of intercellular propagation of metabolites and ROS, most likely by callose deposition (Fig. 2). This helps to keep the carbohydrate level in infected cells high and thereby stimulates the metabolic shift of source cells to a non-assimilatory and carbohydrateconsuming (heterotrophic) metabolism, while assimilation is kept high in adjacent, uninfected cells. The data also suggest that in each cell, the decline in photosynthesis is an irreversible pass-or-fail process, followed by hypersensitive cell death. In our opinion the shift towards a non-assimilatory, carbohydrate consuming metabolic state is at least a prerequisite for, or an integrated part of, hypersensitive reaction and cell death. Defence and hypersensitive reaction do not require light (Mateo et al. 2004). Photosynthesis may even impede or prevent cellular processes required for defence. In our system, we have observed the formation of hypersensitive lesions also in the dark, perhaps even somewhat faster than in light (data not shown). Accumulation of sugars not only fuel carbohydrateconsuming reactions, but also regulates gene expression. High levels of carbohydrates suppress assimilatory genes and stimulate sink-specific genes (for review, see Rolland et al. 2002). In this sense, the initial increase in apoplastic sugars could initiate a self-stimulating feed-forward con-
Figure 8. Metabolic and cellular changes preceding cell wall fortification and cell death during plant defence. Further explanations in the text. © 2005 Blackwell Publishing Ltd, Plant, Cell and Environment, 28, 1421–1435
Plant primary metabolism during defence 1433 trol, by which the metabolic shift toward a sink-type metabolism is further stimulated. It likely explains the biphasic increase of the defence-induced apoplastic invertase. Soluble hexoses also induce the expression of PR proteins and enhance resistance to pathogens (Herbers et al. 1996a, b, 2000). Here, we demonstrate the expression of the sugar-sensitive PR-Q protein (Fig. 1). In this sense, retaining of carbohydrates and accumulation of sugars initiates a shift in the cellular state, both at the level of metabolic regulation and, in the longer term at the level of gene expression. The following scheme (Fig. 8) summarizes the metabolic scenario describe here. Overall, it fits to a number of observations made for other pathosystems, such as, for example for viral infections (Herbers et al. 2000) and to the current understanding of the cellular control of carbohydrate metabolism. However, we introduce the ideas that callose plays a key role in initiating the ‘high sugar state’ and that depression of photosynthesis at the level of photosynthetic electron transport is a prerequisite for the metabolic shift in the cytosol towards respiratory activity and OPPP and, thus, for defence and hypersensitive cell death. We certainly need to experimentally prove some of these ideas, indicated in Fig. 8 with question marks: (1) conclusive evidence for the proposed role of early callose in blocking carbohydrate export; (2) the mechanisms leading to depression of high potential chain activity; (3) a more detailed evaluation of the metabolic processes connecting the decline in photosynthesis and cytosolic processes, in particular OPPP and ROS accumulation; (4) conclusive evidence for the proposed role of ROS in stomatal closure. Also, it has to be evaluated if the scenario described here is specific for the incompatible Phytophthora–tobacco interaction or whether it applies more generally to plant defence reactions.
ACKNOWLEDGMENTS The authors are grateful to Jutta Essmann and Dr Ina Schmitz-Thom for accomplishing the enzyme-activity and sugar determinations. Dr Mark-Aurel Schöttler and Dr Helmut Kirchhoff for assistance with the P700-measurements. Dr Mark-Aurel Schöttler and Dr Ina Schmitz-Thom for critically reading the manuscript. We also thank Prof Dr Uwe Sonnewald (IPK, Gatersleben, Germany) and Prof Dr Thomas Rausch (Ruprecht-Karls-Universität, Heidelberg, Germany) for providing cDNAs.
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