Stem CO2 release under illumination C. Wittmann et al.
Plant, Cell and Environment (2006) 29, 1149–1158
doi: 10.1111/j.1365-3040.2006.01495.x
Stem CO2 release under illumination: corticular photosynthesis, photorespiration or inhibition of mitochondrial respiration? CHRISTIANE WITTMANN1, HARDY PFANZ1, FRANCESCO LORETO2, MAURO CENTRITTO3, FABRIZIO PIETRINI2 & GIORGIO ALESSIO2 1
Institute of Applied Botany, University of Duisburg-Essen, 45117 Essen, Germany, 2CNR – Istituto di Biologia Agroambientale e Forestale and 3CNR – Atmospheric Pollution Institute, 00016 Monterotondo Scalo, Italy
ABSTRACT In illuminated stems and branches, CO2 release is often reduced. Many light-triggered processes are thought to contribute to this reduction, namely photorespiration, corticular photosynthesis or even an inhibition of mitochondrial respiration. In this study, we investigated these processes with the objective to discriminate their influence to the overall reduction of branch CO2 release in the light. CO2 gas-exchange measurements of young birch (Betula pendula Roth.) branches (< 1.5 cm) performed under photorespiratory (20% O2) and non-photorespiratory (< 2%) conditions revealed that photorespiration does not play a pre-dominant role in carbon exchange. This suppression of photorespiration was attributed to the high CO2 concentrations (Ci) within the bark tissues (1544 ± 227 and 618 ± 43 mmol CO2 mol-1 in the dark and in the light, respectively). Changes in xylem CO2 were not likely to explain the observed decrease in stem CO2 release as gasexchange measurements before and after cutting of the branches did not effect CO2 efflux to the atmosphere. Combined fluorescence and gas-exchange measurements provided evidence that the light-dependent reduction in CO2 release can pre-dominantly be attributed to corticular refixation, whereas an inhibition of mitochondrial respiration in the light is unlikely to occur. Corticular photosynthesis was able to refix up to 97% of the CO2 produced by branch respiration, although it rarely led to a positive net photosynthetic rate. Key-words: stem respiration; xylem CO2 concentration.
INTRODUCTION The twigs and branches of most woody plants are able to photosynthesize. In illuminated limbs, the release of CO2 is markedly reduced. It has been shown that this CO2assimilation is performed by bark chlorenchymes (Geurten 1950; Foote & Schaedle 1976a,b; Pilarski 1993; Kharouk et al. 1995; Pfanz et al. 2002). Bark tissues contain Correspondence: Hardy Pfanz. Fax: +49 201 183 4219; e-mail:
[email protected] © 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd
chlorophyll, functional chloroplasts (Pfanz et al. 2002) and active ribulose 1·5-bisphosphate carboxylase/oxygenase (Rubisco) (Schaedle & Brayman 1986), and are therefore competent to photosynthetically refix respiratory CO2. Although net photosynthetic uptake of CO2 is rarely found, corticular refixation of CO2 in young twigs and branches may compensate for 60–90% of the potential respiratory carbon loss and may occasionally exceed CO2 release (Wittmann, Aschan & Pfanz 2001; Pfanz et al. 2002). In most studies, corticular photosynthesis was hence assumed to be equal to the reduction of stem CO2 efflux in the light (Rl) and was calculated as A = |Rd − Rl|,
(1)
where A is the rate of corticular photosynthesis and Rd is stem CO2 efflux in the dark (assumed to be mitochondrial dark respiration). It is not clear in stems and branches, however, if photorespiration within the bark chlorenchymas positively contributes to stem CO2 release under illumination or if (non-photorespiratory) mitochondrial CO2 release is even inhibited by light. Both processes would directly or indirectly effect Rl and thus bias the output of Eqn 1. Furthermore, it could not be excluded that CO2 dissolved in the xylem sap and not generated by local respiring cells could differentially contribute to Rd and Rl. Consequently, all these processes need to be quantified properly to accurately predict the fraction to which corticular photosynthesis contributes to the reduction of stem CO2 release under illumination. In fact, most studies on leaf respiration have reported that the rate of mitochondrial respiration in the light is less than that in darkness (Brooks & Farquhar 1985; Krömer 1995; Atkin et al. 2002), with the degree of inhibition ranging from 16 to 77%. Under illumination, mitochondrial respiration may be down-regulated because photosynthesis supplies sufficient amounts of ATP and NADPH or the enzymes of the tricarboxylic acid cycle are potentially inactivated (Loreto, Velikova & Di Marco 2001; Pinelli & Loreto 2003). Nevertheless, in stems these processes have never been accurately determined. In this study, we therefore combined fluorescence and gas-exchange measurements in young trees of birch (Betula 1149
1150 C. Wittmann et al. pendula Roth.) to investigate the processes that contribute to stem CO2 fluxes under illumination, and, in particular, to ascertain whether the reduction of stem CO2 emission in the light can be attributed in pars or in toto to corticular photosynthesis.
MATERIALS AND METHODS Plant material The experiments were carried out on 6-year-old trees of silver birch (B. pendula Roth.). The trees were grown outside in 20 L plastic containers under sufficient nutrition (Einheitserde Typ T, Balster, Germany) and water supply, realized by periodic fertilization with Osmocote (Bayer, Leverkusen, Germany) and daily irrigation to avoid water and nutrient stresses. For fluorescence and gas-exchange measurements, ten trees were transferred whole to the laboratory and for each tree, all parameters were measured at the middle of the length of one to two 1-year-old stems (n = 10–20). In all cases, measurements were performed on stems from the outer sun-crown at a height of about 1.5 m.
Epifluorescence microscopy Cross-sections of current- and 1-year-old birch twigs were visualized with an epifluorescence microscope (Olympus GmbH, Hamburg, Germany) equipped with a blue exitation filter (BP 450–490) and an emission filter (LP 520) and linked to a computer. Images were captured with an digital Insight camera (Visitron Systems Inside QE, Puchheim, Germany), and image contrast was enhanced using diagnostic imaging advanced SPOT software (SPOT visitronsystems, Puchheim, Germany). Chlorophyllous cells were identified by their bright red fluorescence, which is characteristic of chlorophyll.
CO2/H2O gas-exchange and fluorescence measurements For fluorescence and gas-exchange measurements, intact stems with a diameter < 1.5 cm were used. A 8-cm-long portion of stem was enclosed in a plastic cuvette allowing simultaneous measurements of CO2 and H2O exchange by infrared gas analysis and chlorophyll fluorescence. The used system allows to operate with full control of O2 and CO2 concentrations, incident light intensity, air temperature and relative humidity inside the cuvette. Stem temperature was measured with a thermocouple attached to the outer stem surface. Light was supplied by a Osram Power Star HQI-R lamp (OSRAM GmbH, München, Germany). The stem segment was exposed to a flow of 400 mL min−1 of synthetic air (N2, O2 and CO2) formed and adjusted with mass-flow controllers (Brooks, Veenendaal, The Netherlands). The synthetic air mixture was humidified by bubbling it in water and condensing part of the humidity in a water bath set at a temperature lower than the stem temperature. CO2 and
H2O exchanges between the stem and the air were measured with a differential infrared gas analyser (Li 6262; Li-Cor, Lincoln, NE, USA). The CO2 release in the dark (Rd) was measured after 45 min of exposure to darkness. Subsequently, CO2 release in the light (Rl) was determined after a 45 min period of light exposure to 600 µmol photons m−2 s−1. All measurements were made under constant microclimatic conditions (relative humidity of 45 to 50% and constant temperatures of 20 or 30 °C), under ambient CO2 (350 µmol mol−1) and photorespiratory (20% O2) or non-photorespiratory (2% O2) conditions. Because externally supplied gases may penetrate the outer bark slowly (Shepard 1970), the time of exposure to different gas concentrations was set from 45 to 120 min until a steady-state exchange of CO2 and fluorescence yield was measured. A polyfurcated optic fibre of a modulated fluorometer (MiniPAM, Walz, Effeltrich, Germany) was attached to the stem to measure fluorescence under actinic light (600 µmol photons m−2 s−1) and pulses of saturating light (> 10 000 µmol photons m−2 s−1).
Total periderm conductance to water vapour and CO2 According to Campbell & Norman (1998) and Cernusak & Marshall (2000), total water vapour conductance of the stem surface/periderm was calculated as gtw =
E
(wi − wa )
,
(2)
where gtw, is the total conductance to water vapour, E is the peridermal or stem transpiration rate (µmol H2O m−2 s−1), wi is the molar concentration of water vapour within the stem and wa is the molar concentration of water vapour of the atmosphere (µmol H2O mol air−1). The vapour conductance of air beneath the stem surface was assumed to be saturated; that of the atmosphere was calculated from the relative humidity and temperature data. The transpiration rate was determined from the H2O exchange measurements. The total periderm conductance for CO2 was determined for each stem segment as gtc =
gtw , 1.56
(3)
where gtc is the total conductance to CO2, gtw is the total conductance to water vapour and 1.56 is the ratio of the diffusivities of water and CO2 in air (Campbell & Norman 1998). It has conclusively been demonstrated that the bark and lenticel intercellular spaces are not filled with water and form a continuous gas phase (Hook & Brown 1972; Hook, Brown & Wetmore 1972; Schönherr & Ziegler 1980; Langenfeld-Heyser et al. 1996; Grosse 1997; LangenfeldHeyser 1997). Thus, an unhindered gas flow can occur not only in leaves, but also in the intercellular system of the bark. Consequently, we used for the calculation the same ratio of the diffusivities of water and CO2 that are generally used for leaves (1.56)(Campbell & Norman 1998).
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
Stem CO2 release under illumination 1151
Calculation of corticular CO2 concentration and electron transport rate by fluorescence and gas-exchange According to Fick’s law of diffusion, the corticular CO2 concentration (cort. Ci) is most simply calculated as cort. C i = Ca −
AN , gtc
(4)
where AN is the net CO2 efflux from the stem segment, gtc is the total conductance of the stem surface to CO2, Ca is the concentration of CO2 in the atmosphere and cort. Ci is the concentration of CO2 in the intercellular system of the stem cortex (Cernusak & Marshall 2000). The electron transport rate (ETR) can be derived from fluorescence measurements (ETRf) as the product of photosystem II (PSII) quantum yield (measured by the parameter ∆F/Fm’, cfr Genty, Briantais & Baker 1989) and quantum flux density of the incident photosynthetically active radiation (PAR, µmol m−2 s−1), assuming an equal distribution between the two photosystems and, thus only 50% of the absorbed light impinges on PSII (denoted by 0.5) ETRf = ∆F/Fm’* (PAR * (Tp/100)) * 0.5.
(5)
The factor (Tp/100) takes into account the amount of light transmitted by the periderm. The value of Tp, calculated for 1-year-old birch twigs, was 25.64 ± 1.76 (400–700 nm). That means that approximately 26% of the incident photosynthetically active quanta reach the bark chlorenchyme. Thus, ETRf = ∆F/Fm’ * (PAR * 0.26) * 0.5.
(6)
The ETR can be also calculated from gas-exchange measurements (ETRg) by multiplying the photosynthetic rate by four (Loreto et al. 1994) under the assumption that a minimum of four electrons are required to fix one mole of CO2 (or that four electrons are generated for each molecule of O2 released) when photorespiration is suppressed. Photosynthesis was assumed to be equal to the reduction of respiration observed in illuminated leaves with respect to darkened leaves (Eqn 1). Thus, ETRg = A * 4 = |Rd − Rl| * 4
(7)
If we assume that, under non-photorespiratory conditions, photosynthesis is the only metabolic sink for electrons, the measured values of ETR should give identical results (ETRg = ETRf).
Temperature response measurements To determine the temperature response of leaf and stem chlorophyll fluorescence, whole trees were transferred to the laboratory and were placed in a climate chamber (PVP GmbH, Willich, Germany), which allows full control of temperature, relative humidity and light intensity. All fluorescence measurements were made under ambient CO2 and constant microclimatic conditions (relative humidity: 40– 45%; ∼600 µmol photons m−2 s−1) by means of a pulse-
amplitude-modulated fluorimeter (PAM-2000; Walz), equipped with a 2030-B leaf-clip holder, featuring an integrated microquantum sensor and a thermocouple. The 0.8 s saturation pulse intensity was set to approximately 10 000 µmol m−2 s−1; an intensity that was found to be always saturating. To accurately determine the fluorescence signal of a cylindrical stem, we attached a non-fluorescing black paper with a rectangular 3 × 10 mm window on the clip. During the measurements, this window was orientated along the stem axis, thus only photon exchange between the plane centre of the stem and the instrument is considered. Outer bark temperature of the investigated stems was measured by the thermocouple of the clip holder directly attached to the stem surface; all temperatures reported are thus stem surface temperatures (± 0.5 °C). All measurements were made as follows: chamber temperatures were set to a constant value and the trees were left to temperature-adapt for at least 60 min. Then the effective quantum yield of PSII was measured on two to three intact leaves and 1-year-old stems of each tree (n = 5) under an actinic light intensity of 600 µmol photons m−2 s−1. Thereafter, the trees were exposed to a new temperature level. Generally, the temperature was altered sequentially, from 5 to 43 °C. In parallel with the measurements of chlorophyll fluorescence, we determined Rd. All measurements were performed after 60 min dark adaptation under a constant relative humidity (45–50%) and a controlled CO2-supply (350 µmol mol−1) using an infrared gas analyser (LI-6400).
O2 gas-exchange measurements Respiration in Rd and Rl was determined also from measurements of oxygen gas exchange in detached bark samples, using the Hansatech LD2 liquid-phase oxygen electrode system (Hansatech, King’s Lynn, Norfolk, UK) designed by Delieu & Walker (1981). All measurements were made at a thermostatically controlled (Bachofer, Reutlingen, Germany) temperature of 20 °C and a constant, optimal bicarbonate concentration (3 mM KHCO3 and 1 µL carbonic anhydrase). The optimal bicarbonate concentration was determined from A/Ca curves (data not shown). Illumination of the chamber was provided by a 50 W halogen dichroic lamp (Osram), with the light passing through a water bath to remove infrared heat. Neutraldensity filters (Balzers AG, Balzers, Liechtenstein) were used at the surface of the chambers to attenuate the light to the desired levels. Just prior to use, 1 cm2 fresh bark was peeled from the stem and carefully infiltrated with incubation medium. The incubation medium consisted of Mes (morpholino-ethane-sulfonic acid), KCl, MgSO4, Ca(NO3)2, KHCO3 of 50, 4, 1, 1 and 3 mM, respectively (+ 1 µL carbonic anhyhrase; activity: 2000 U mg−1), buffered at pH 6. The pieces were placed in a 50 mL plastic syringe with 20 mL incubation medium (assay buffer) with all the air removed. A gentle vacuum was created by withdrawing the plunger. During the vacuum, the syringe was agitated to dislodge any gas bubbles from the surface of the tissues. The vacuum was released and a slight positive
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
1152 C. Wittmann et al. pressure was applied (Pfanz & Dietz 1987). The bark pieces were suspended from the cuvette stopper by means of a small plastic-coated nichrome wire which was threaded through the injection hole and oriented to the light source. Calibrations and calculations were described by Walker (1987). All measurements were made as follows: (1) samples were left to dark-adapt for at least 30 min until a steady rate of dark respiration was attained; (2) in the next step, the samples were pre-illuminated for 5–10 min (at 150 µmol photons m−2 s−1) and then left to adapt to an initial irradiation of around 25 µmol photons m−2 s−1 until a steady-state rate of O2 evolution was reached; and (3) thereafter, the samples were exposed to a new light step. Working with excised tissues could alter O2 gas exchange by inducing wound respiration (e.g. Sprugel & Benecke 1991). To determine whether excision influenced our measurements, we left isolated birch bark samples for 90 min in the dark recording the O2 gas exchange. Oxygen consumption increased transiently to reach a steady-state rate within 30–40 min. In the following 50–60 min, no changes in oxygen exchange rate were found, indicating that excision had not greatly altered respiration.
Statistics For statistical data analysis, the SigmaPlot 2001 software (SPSS inc., Chicago, IL, USA) was used. Significance of differences between sets of data were checked by Student’s t-tests. To visualize and plot the curve that best describes the shape and behaviour of our data (‘curve fitting’), the ‘regression wizard’ of the programme was used. For the CO2- and O2-exchange versus PAR curves, we found that an exponential equation (‘exponential rise to maximum’) most closely describes, or fits, the data. The cort. Ci versus PAR curves were treated as exponential decays and the Rd versus diameter curves as linears. The coefficient of determination, a measure of how well the regression model describes the data, is given in the description of the figures. The SigmaPlot curve fitter uses the Marquardt–Levenberg algorithm to find the coefficients (parameters) of the independent variable(s) that give the ‘best fit’ between the equation and the data. This algorithm seeks the values of the parameters that minimize the sum of the squared differences between the values of the observed and predicted values of the dependent variable. For details about curvefitting algorithms see Marquardt (1963).
RESULTS AND DISCUSSION Illumination of young branches of birch typically decreases the CO2 efflux to the atmosphere. In the dark, respiratory CO2 is formed and released to the atmosphere. The decrease of this efflux upon illumination may be caused and influenced by several independent or interdependent factors: 1 Photorespiration might be a factor contributing to the CO2 budget under illumination.
2 Corticular photosynthesis refixes CO2 derived from mitochondrial respiration, thus reducing the stem CO2 efflux. 3 Mitochondrial respiration could be reduced in the light, as often observed in leaves. 4 CO2 dissolved in the xylem sap and not generated locally by respiring cells could differentially contribute to Rd and Rl. Whether or not these processes act on stem CO2 gas exchange is the topic of this article and discussed below.
The influence of photorespiration on stem CO2 efflux Photorespiration within the bark chlorenchymes may positively contribute to stem CO2 release under illumination. Several authors already hypothesized that photorespiration is likely to be restricted in the cortex chlorenchymes as stem-internal CO2 concentrations are assumed to be rather high and O2 concentrations concomitantly lower (e.g. Schaedle 1975; Foote & Schaedle 1976a, Comstock & Ehleringer 1990; Levy & Jarvis 1998; Pfanz et al. 2002). In fact, published values for xylem CO2 concentrations ranged from 1 to 26% (see McDougal & Working 1933; Jensen 1969; Carrodus & Triffett 1975), while CO2 concentrations in bark tissues are thought to be around 1000 to 1700 µmol mol−1 (Cernusak & Marshall 2000). Our calculations of cort. Ci using data derived from CO2 gas-exchange measurements confirm these results. In intact birch branches (< 1.5 cm diameter), the mean corticular CO2 was 1548 ± 227 and 618 ± 43 µmol CO2 mol−1 in the dark and light, respectively (after a 45 min illumination at 600 µmol photons m−2 s−1). In all cases, cort. Ci were substantially higher in the dark and decreased gradually with increasing light intensity (Fig. 1b), thus mirroring the onset of lightdependent processes that decrease the net efflux of CO2 out of the stems (Fig. 1a). For leaves, however, published Ci-values are around 180 to 240 µmol CO2 mol−1 for C3 or C4 plants, respectively (Von Willert et al. 1995). Estimates of cort. Ci were thus up to six times higher. Consequently, a suppression of photorespiration due to high cort. Ci cannot be excluded. To further investigate whether photorespiration contributes to the gas exchange of birch branches, we performed experiments under normal, photorespiratory (20% O2) and nonphotorespiratory (< 2% O2) conditions. Again, Rl was found to be lower than Rd, but both rates were not sensitive to differences in O2 concentrations and, consequently, |Rd − Rl| (corticular photosynthesis) was also not affected by O2 concentration (Fig. 2). As the oxygen concentration strongly determines the ratio between photosynthesis and photorespiration, these results hint to the fact that photorespiration does not play a pre-dominant role in CO2 exchange of illuminated birch stems. The relative insensitivity of corticular photosynthesis may also be explained by a C4-like pathway of corticular photosynthesis. Hibberd & Quick (2002) hypothesized that C4 photosynthesis first developed in bundled sheath cells of
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
Stem CO2 release under illumination 1153
0.0
Rd
(a)
–0.6 –0.8 –1.0
(mmol CO2 m–2 s–1)
CO2 exchange rate
–2 –1
(mmol CO2 m s )
CO2 exchange rate
–0.2 –0.4
Rl
2.0
ns
1.5 1.0 0.5 0.0 –0.5
ns
–1.0 –1.5
–1.2
0
200
400
600
800
1000
(b)
1800 –1
(mmol CO2 mol )
–2.0
200 mbar O2
ns
20 mbar O2
–2.5
2000 Corticular CO2 concentration
|Rd – Rl|
Figure 2. CO2 release of one-year-old twigs of Betula pendula in the dark (Rd) and in the light (Rl; after 45 min under illumination with 600 µmol photons m−2 s−1). The absolute value of the difference (|Rd − Rl|) is generally interpreted to be a measure of corticular photosynthesis. Measurements were performed at 200 and 20 mbar O2 under constant climatic conditions (20 °C, 45–50% relative humidity) and at controlled CO2-supply (350 µmol mol−1). Data are means ± SD (n = 10), ns = differences are statistically not significant (P > 0.05).
1600 1400 1200 1000 800 600 400 0
200
400
600
800
1000
PAR (mmol photons m–2 s–1)
Figure 1. (a) Light response of stem CO2-efflux as measured in one-year-old intact twig segments of Betula pendula. Measurements were performed under constant climatic conditions (20 °C, 45–50% relative humidity) and a controlled CO2-supply (350 µmol mol−1) using a CO2 porometer (LI-6400). (b) Light response of corticular CO2 concentration (cort. Ci) as calculated from gas-exchange measurements (means with ± SD, n = 10). PAR, photosynthetically active radiation.
pression of photorespiration as well as a shift of the photosynthetic optimum to higher temperatures (Acock & Allen 1985; Long 1991; Bowes 1993). We assume that both rising temperature and high cort. Ci influence corticular photosynthesis through their direct effects at the level of primary carboxylation. In addition, increased temperature favours both the solubility of CO2 and the specificity of Rubisco for CO2 (Long 1991). An increased cort. Ci favours ribulose bisphosphate carboxylation by inhibition of acceptor oxygenation. Table 1. Measurements of light-dependent reduction of CO2
stems and petioles of C3 plants and later became common in leaves. They measured high activities of the decarboxylating enzymes involved in C4 photosynthesis in extracts from vascular strands taken from C3 plants. In several deciduous trees, phosphoenolpyruvate (PEP)-carboxylase was found at around 10 to 23 times levels higher than in leaves of C3 plants (Höll 1973, 1974; Berveiller & Damesin 2005). PEP-carboxylase, an enzyme present in nearly all cells, is able to fix dissolved CO2 (HCO3–) in quite high amounts (Müller, Baier & Kaiser 1991), but has no affinity for O2. Furthermore, we detected a temperature-dependent stimulation of corticular photosynthesis (Table 1) and photochemical yield in illuminated stems (Fig. 3). The latter parameter showed a clear difference with respect to birch leaves, with a temperature maximum between 32 and 36 °C in stems and 26 and 30 °C in leaves. Thus, compared to most C3 plants, where the photosynthetic optimum is in the range of 25–30 °C (Larcher 2001), optimal corticular photosynthesis occurs at higher temperatures. In leaves of C3-plants, elevated CO2 concentrations have shown to cause a sup-
release (|Rd − Rl|), photosynthetic electron transport rate measured by gas-exchange and fluorescence and corticular refixation (%) in Betula stems. Parameters
20 °C
30 °C
Stem CO2 efflux (µmol m−2 s−1) Rd Rl |Rd − Rl|a Corticular refixation (% of Rd)
−1.46 ± 0.10a −0.07 ± 0.10a 1.39 ± 0.10a 97.0 ± 5.0a
−2.28 ± 0.10b −0.48 ± 0.10b 1.82 ± 0.20b 75.0 ± 4.0b
Electron transport rate measured by (µmol m−2 s−1) ETRg (CO2 gas-exchange) 5.56 ± 0.40a ETRg (O2 gas-exchange) 13.8 ± 1.1a ETRf (fluorescence) 13.4 ± 0.2a
7.20 ± 0.50b 25.5 ± 1.9b 26.9 ± 0.3b
Letters indicate temperature-dependent differences statistically significant at P < 0.05. Measurements were performed at air temperature of 20 and 30 °C and 600 µmol photons m−2 s−1. Means ± SE (n = 10) are reported. a A = |Rd − Rl| = |−1.46 + 0.07| = 1.39; ETRg = 4 * A = 5.56. Rd, stem CO2 efflux in the dark; Rl, stem CO2 efflux in the light; ETRg, electron transport rate by gas-exchange; ETRf, electron transport rate by fluorescence; A, rate of corticular photosynthesis.
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
1154 C. Wittmann et al. the periderm, the actual light intensity impinging on the branch chlorenchyme is substantially lower than the incident light at the outer branch surface. Correction of the light intensity according to the peridermal light transmittance led to an clear increase in steepness of the light response curve (Fig. 4), which can be attributed to the fact that bark chlorenchyme display many characteristics compatible with a shade-adaptation syndrome (see also Schmidt, Batic & Pfanz 2000; Wittmann et al. 2001; Pfanz et al. 2002; Damesin 2003; Manetas 2004). However, light saturation of photosynthesis was reached also at higher light intensity and was higher in peeled bark tissues than in intact bark tissues. Thus, we can conclude that the periderm is an effective barrier to radial diffusion of CO2 from bark tissues. Peeling of the bark tissues altered or removed these barriers, and increased gas diffusion as well as light availability of the underlying bark tissues.
Stem Leaf 0.5
0.4
0.3
0.2 6 °C 0.1
0.0
0
10
20
30
40
50
Surface temperature (°C) Figure 3. Temperature response of photosystem II (PSII)efficiency of leaves and intact young stems (< 2-cm diameter) of Betula pendula. Measurements were performed at 600 µmol photons m−2 s−1, a constant CO2/O2 supply (350 µmol CO2 mol−1; 200 mbar O2) and a constant relative humidity (45–50). Data are means ± SE (n = 10–20).
Stem-internal CO2-recycling corticular photosynthesis When experiments were performed with intact branches, corticular photosynthesis expressed as a percentage of dark CO2 efflux reduced up to 97% of Rd (Table 1), but did not lead to a positive net carbon uptake. Using peeled bark tissues in aqueous systems led to different results. Removal of the periderm resulted in a substantial increase in lightdependent O2 evolution, which even led to a positive net photosynthetic rate of up to 2 µmol O2 m−2 s−1 (Fig. 4). A fact that we attribute mainly to the low conductance of the periderm (for water: 1.05 ± 0.25 mmol m−2 s−1, n = 15; for CO2: 0.50 ± 0.16 mmol m−2 s−1, n = 15). Cernusak & Marshall (2000) found a comparable bark conductance to water vapour of 1.03 mmol m−2 s−1 and a corresponding CO2 conductance of 0.68 mmol m−2 s−1 in 4-year-old branches of Pinus monticola. However, stem periderms are not fully impermeable. They show measurable permeances for water and gases (Geurten 1950; Schönherr & Ziegler 1980; Buchel & Grosse 1990; Langenfeld-Heyser et al. 1996, Langenfeld-Heyser 1997, Groh, Hübner & Lendzian 2002), mainly accomplished by means of lenticels which cover between 2 and 10% of the stem area (in leaves, stomata occupy between 0.2 and 2.0% of the surface [Meidner & Mansfield 1968]). However, we assume that the observed difference in light-dependent O2 evolution can be explained by: (1) a better gas permeability; and (2) an improved light availability of the underlying bark tissues after periderm removal. Due to the optical properties of
Inhibition of mitochondrial respiration under illumination and its influence on stem CO2 efflux In the absence of photorespiration and under optimal, nonstress conditions, photosynthesis is the only metabolic sink for electrons, and the photosynthetic ETR can be estimated either from ETRg or from ETRf (Loreto et al. 1994). Thus, under non-photorespiratory conditions, measurements of ETR should yield the same value (ETRg = ETRf). Any discrepancy from this equation would indicate that the difference between Rd and Rl is not solely due to corticular refixation. Furthermore, a light-dependent inhibition of mitochondrial respiration, similar to that often detected in leaves (first by Kok 1948) could be responsible, in pars or in toto, for |Rd − Rl| (and ETRf − ETRg) differences. Under this assumption, we would expect that ETRf would be lower than ETRg, as an inhibition of mitochondrial respi-
3 O2 exchange rate ( mmol O2 m–2 s–1)
Photochemical PSII efficiency (yield)
0.6
2 1 0 –1 –2
Peeled, R2 = 0.91 Not peeled, R2 = 0.84 Not peeled, corrected PAR
–3 –4 0
200
400
600
800
1000
1200
1400
PAR ( mmol photons m–2 s–1)
Figure 4. Light response of net photosynthesis of isolated bark chlorenchyme as measured in an oxygen electrode under optimum conditions. Means ± SE (n = 10). PAR, photosynthetically active radiation.
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
Stem CO2 release under illumination 1155
sc cp
sc
ph
cp
xy
ph xy mp
(a)
(b)
ration would not yield photosynthetic electron transport. However, in our experiments ETRf is clearly higher than ETRg (Table 1). On one hand, this excludes a substantial contribution of mitochondrial respiration in the determination of the gradient between Rd and Rl. On the other hand, it could also indicate that photorespiration is not fully suppressed in stems and thus contributes to ETRf (or that ETRg is lowered by the high diffusive resistance of the periderm). Yet, measurements under low O2 (Fig. 2) did not indicate that photorespiration is an active process in stems. The calculation of ETRg made on the basis of O2 gas-exchange measurements of peeled branches leads to an ETRg being very similar to ETRf (Table 1). This is in accordance to our assumption that in the absence of photorespiration, photosynthesis is the only metabolic sink for electrons. In this case, inhibition of mitochondrial respiration in the light is unlikely to occur. We speculate that fluorescence only detects a superficial layer of chloroplasts (Fig. 5), and that this matches the layer of photosynthetic tissue exposed to gas exchange after periderm removal, but not in intact stems, thus explaining the difference between ETRf and ETRg in the latter specimen.
The potential influence of xylem sap flow As recently published, CO2 moving in the transpiration stream may alter xylem CO2 and affect the rate of CO2 diffusion from woody tissues to the atmosphere (Teskey & McGuire 2002, 2005; Gansert 2003). Yet, xylem CO2 reaching assimilatory tissues can be reassimilated, thereby modifying assimilation and respiration rates calculated solely –1000
(a)
R 2 = 0.02 (P = 0.5728)
Rd (m mol CO2 m–3 s–1)
Rd (m mol CO2 m–2 s–1)
–3.5 –3.0 –2.5 –2.0 –1.5 –1.0 –0.5 0.0 2
4
6
8
10
Diameter (mm)
12
14
Figure 5. Cross-sections of (a) a current year and (b) one-year-old stem of Betula pendula observed by epifluorescence microscopy. Red chlorophyll fluorescence was excited by blue light. cp, cortex parenchyma; sc, sclerenchyma; ph, phloem; xy, xylem; mp, medullary parenchyma.
on the basis of gas-exchange measurements (e.g. Negisi 1974; Martin, Teskey & Dougherty 1994; Hibberd & Quick 2002). When CO2 efflux before and after cutting of the branches was measured (for 4 h), no significant differences were found (data not shown). Similar results have also been reported by Gansert (2003) who found that stem CO2 efflux rates of birch were not affected by a gradual decrease of sap flow. The data presented in Fig. 6 are in good agreement with these results and further rule out xylem CO2 as a major control of carbon efflux in the branches under investigation (< 2 cm diameter). If the living tissues (as the source of respiratory CO2) are assumed to be concentrated along the perimeter of the cross-section of the stem (Kinerson 1975; Linder & Troeng 1981; Lavigne 1988), surface area-related respiration should be expected to be scale-invariant, while a volume-based measure would be proportional to the reciprocal of stem diameter (d−1). We found a positive linear relationship between d−1 and Rd per unit volume, but no relationship between CO2 efflux and stem area (Fig. 6). Branch CO2 efflux to the atmosphere is thus generated mainly from the local sources (cambial, cortical and phloem cells), while the contribution of xylem CO2 seems to be negligible in our experiments. Working with young limbs (1 to 2-years-old specimens), we have to point out that the high surface to volume ratio minimizes the contribution of xylem CO2, and live sapwood respiration rates are small in comparison to phloem/cambial respiration. In older and larger trees where sapwood makes up more than 80% of the total stem volume, the contribution of CO2 from xylem vessels is clearly larger (McGuire & Teskey 2004; Teskey & McGuire 2005).
R 2 = 0.74 (P < 0.0001)
(b)
–800 –600 –400
Figure 6. Dark stem CO2 efflux (Rd) of
–200 0 0.00 0.05 0.10 0.15 0.20 0.25 0.30 Diameter–1 (mm–1)
young birch stems of a range of diameters (d), expressed on a surface area (a) and volume area (b) basis. Measurements were performed in the dark under constant climatic conditions (20 °C, 45– 50% relative humidity) and a controlled CO2-supply (350 µmol mol−1).
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
1156 C. Wittmann et al. However, even if an influence of xylem sap flow on stem CO2 efflux is expected, it should be of minor importance for the observed phenomenon. Because, if illumination would induce an increase in temperature within the xylem vessels of the wood, CO2 degassing from the vessels should increase; temperature profoundly changes the holding capacity of water, thus a 5 °C increase could drive 12% of the CO2 out of solution (Rakonczay 1997). Consequently, a higher efflux of CO2 from the branches into the atmosphere would occur. Yet, the opposite is observed. The CO2 efflux is substantially reduced under illumination (Fig. 1). Furthermore, to avoid any possible changes in tissue temperature, the surface temperatures of the branches within the measuring cuvettes were kept constant. Thus, a light-induced shift in tissue temperature and its probable influence on xylem sap flow cannot explain the observed decrease in stem CO2 release under illumination.
CONCLUSIONS The precise measurement and evaluation of gas exchange within twigs and branches and with the atmosphere is very crucial for understanding corticular metabolism. Yet, data are not easily to be compared as all the techniques used have their specific restrictions. Under optimum conditions, corticular photosynthesis is able to refix up to 97% of the CO2 produced by the respiration of young branches of birch (Table 1). Intact branches rarely reveal a positive net photosynthetic rate. Yet, O2 gas-exchange measurements using peeled stems clearly prove that the photosynthetic capacity of the cortex is much higher, but is masked by peridermal resistances. Removal of the periderm clearly increased ‘measurable’ corticular photosynthesis (up to 2 µmol O2 m−2 s−1). Corticular photosynthesis determined on the basis of gas-exchange measurements of intact branches are an integration over the whole stem organ, while measurements using fluorescence signal most probably describe the reactions of the topmost stem layers (cambial, phloem and bark tissues). Yet, it seems as if exactly these topmost layers are the responsible tissue fractions measured in gas exchange. There is some evidence that photorespiration does not occur in birch stems and that mitochondrial respiration seems to be fairly similar in the light and in the dark. Photorespiratory activity is probably inhibited by the high cort. Ci. In leaves of C3-plants, elevated CO2 concentrations have shown to cause a suppression of photorespiration as well as a shift of the photosynthetic optimum to higher temperatures. The high photosynthetic temperature optimum found in young birch branches hints to a similar effect in the bark tissues of birch stems, probably due to modifications that high cort. Ci impose on the temperature response of corticular photosynthesis. Both increasing temperature and higher cort. Ci directly influence corticular photosynthesis through their direct effects at the level of primary carboxylation.
ACKNOWLEDGMENTS This work was partly performed within the Cost Action 627. We would like to thank Alfred Lenk, Gudrun Friesewinkel, Sabine Kühr and Christa Kosch for their technical assistance. Special thanks also to Dipl. Umweltwiss. Regina Wegner for her comments on the draft manuscript.
REFERENCES Acock B. & Allen L.H. (1985) Crop responses to elevated carbon dioxide concentrations. In Direct Effects of Increasing Carbon Dioxide on Vegetation (eds B.R. Strain & J.D. Cure), pp. 53–97. United States Departement of Energy, Washington, DC, USA. Atkin O.K., Evans J.R., Ball M.C., Lambers H. & Pons T.L. (2002) Leaf respiration of snow gum in the light and dark: interactions between temperature and irradiance. Plant Physiology 122, 915– 923. Berveiller D. & Damesin C. (2005) Abstracts of the 10th International Meeting of the Working Group of Experimental Ecology within the German Society of Ecology. AKOE 2005, 27. Bowes G. (1993) Facing the inevitable: plants and increasing atmospheric CO2. Annual Review of Plant Physiology and Plant Molecular Biology 44, 309–332. Brooks A. & Farquhar G.D. (1985) Effect of temperature on the CO2-O2 specificity of ribulose-1,5-biphosphate carboxylase/oxygenase and the rate of respiration in the light: estimates from gas exchange measurements on spinach. Planta 165, 397–406. Buchel H.B. & Grosse W. (1990) Localization of the porous partition responsible for pressurized gas transport in Alnus glutinosa (L.). Tree Physiology 6, 247–256. Campbell G.S. & Norman J.M. (1998) An Introduction to Environmental Biophysics, 2nd edn. Springer Verlag, Heidelberg, Germany. Carrodus B.B. & Triffett A.C.K. (1975) Analysis of respiratory gases in woody stems by mass spectrometry. New Phytologist 74, 243–246. Cernusak L.A. & Marshall J.D. (2000) Photosynthetic refixation in branches of western white pine. Functional Ecology 14, 300– 311. Comstock J.P. & Ehleringer J.R. (1990) Effects of variations in leaf size on morphology and photosynthetic rate of twigs. Functional Ecology 4, 209–221. Damesin C. (2003) Respiration and photosynthetic characteristics of current-year stems of Fagus sylvatica: from the seasonal pattern to annual balance. New Phytologist 158, 465–475. Delieu T. & Walker D.A. (1981) Polarographic measurement of photosynthetic O2 evolution by leaf discs. New Phytologist 89, 165–175. Foote K.C. & Schaedle M. (1976a) Physiological characteristics of photosynthesis and respiration in stems of Populus tremuloides Michx. Plant Physiology 58, 91–94. Foote K.C. & Schaedle M. (1976b) Diurnal and seasonal patterns of photosynthesis and respiration by stems of Populus tremuloides Michx. Plant Physiology 58, 651–655. Gansert D. (2003) Xylem sap flow as a major pathway for oxygen supply to the sapwood of birch (Betula pubescens Her.). Plant, Cell and Environment 26, 1803–1814. Genty B., Briantais J.M. & Baker N.R. (1989) The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence. Biochimica et Biophysica Acta 990, 87–92. Geurten T. (1950) Untersuchungen über den Gaswechsel von Baumrinden. Forstwissenschaftliches Centralblatt 69, 704–743.
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
Stem CO2 release under illumination 1157 Groh B., Hübner C. & Lendzian K.J. (2002) Water and oxygen permeance of phellems isolated from trees: the role of waxes and lenticels. Planta 215, 794–801. Grosse W. (1997) Gas transport in trees. In Trees-Contributions to Modern Tree Physiology (eds H. Rennenberg, W. Eschrich & H. Ziegler), pp. 57–74. Backhuys Publishers, Leiden, The Netherlands. Hibberd J.M. & Quick W.P. (2002) Characteristics of C4 photosynthesis in stems and petioles of C3 flowering plants. Nature 415, 451–454. Höll W. (1973) Enzyme activities in wood tissue as affected by different methods of homogenizing. Holzforschung 27, 145–147. Höll W. (1974) Dark CO2 fixation by cell-free preparations of wood of Robinia pseudoacacia. Canadian Journal of Botany 52, 727–734. Hook D.D. & Brown C.L. (1972) Permeability of the cambium to air in trees of wet habitats. Botanical Gazette 133, 304–310. Hook D.D., Brown C.L. & Wetmore R.H. (1972) Aeration in trees. Botanical Gazette 133, 443–454. Jensen K.F. (1969) Oxygen and carbon dioxide concentrations in sound and decaying red oak trees. Forest Science 15, 246–251. Kharouk V.L., Middleton E.M., Spencer S.L., Rock B.N. & Williams D.L. (1995) Aspen bark photosynthesis and its significance to remote sensing and carbon budget estimates in the boreal ecosystem. Water, Air and Soil Pollution 82, 483–497. Kinerson R.S. (1975) Relationship between plant surface area and respiration in loblolly pine. Journal of Applied Ecology 12, 965– 971. Kok B. (1948) A critical consideration of the quantum yield of Chlorella-photosynthesis. Enzymologia 13, 1–56. Krömer S. (1995) Respiration during photosynthesis. Annual Review of Plant Physiology and Plant Molecular Biology 46, 45– 70. Langenfeld-Heyser R. (1997) Physiological functions of lenticels. In Trees-Contribution to Modern Tree Physiology (eds H. Rennenberg, W. Eschrich & H. Ziegler), pp. 43–56. Backhuys Publishers, Leiden, The Netherlands. Langenfeld-Heyser R., Schella B., Buschmann K. & Speck F. (1996) Microautoradiographic detection of CO2 fixation in lenticel chlorenchyme of young Fraxinus excelsior L. stems in early spring. Trees 10, 255–260. Larcher W. (2001) Ökophysiologie der Pflanzen: Leben, Leistung und Stressbewältigung der Pflanzen in Ihrer Umwelt, 5th edn. Ulmer, Stuttgart, Germany. Lavigne M.B. (1988) Stem growth and respiration of young balsam fir trees in thinned and unthinned stands. Canadian Journal of Forest Research 18, 483–489. Levy P.E. & Jarvis P.G. (1998) Stem CO2 fluxes in two Sahelian shrub species (Guiera senegalensis and Combretum micanthum). Functional Ecology 12, 107–116. Linder S. & Troeng E. (1981) The seasonal variation in stem and course root respiration of a 20-year-old Scots pine (Pinus sylvestris L.). Miteilungen der Forstlichen Versuchsanstalt 142, 125–139. Long S.P. (1991) Modification of the response of photosynthetic productivity to rising temperature by atmospheric CO2 concentrations: has its importance been underestimated? Plant, Cell and Environment 14, 729–739. Loreto F., Di Marco G., Tricoli D. & Sharkey T.D. (1994) Measurements of mesophyll conductance, photosynthetic electron transport and alternative electron sinks of field grown wheat leaves. Photosynthesis Research 41, 397–403. Loreto F., Velikova V. & Di Marco G. (2001) Respiration in the light measured by 12CO2 emission in 13CO2 atmosphere in maize leaves. Austrian Journal of Plant Physiology 28, 1103–1108. McDougal D.T. & Working E.B. (1933) The Pneumatic System of
Plants, Especially Trees, Publication 441. Carnegie Institution of Washington, Washington, DC, USA. McGuire M.A. & Teskey R.O. (2004) Estimating stem respiration in trees using a mass balance approach that accounts for internal and external fluxes of CO2. Tree Physiology 24, 571–578. Manetas Y. (2004) Probing corticular photosynthesis through in vivo chlorophyll fluorescence measurements: evidence that high internal CO2 levels suppress electron flow and increase the risk of photoinhibition. Physiologia Plantarum 120, 509–517. Marquardt D.W. (1963) An algorithm for least squares estimation of parameters. Journal of the Society of Industrial and Applied Mathematics 11, 431–441. Martin T.A., Teskey R.O. & Dougherty P.M. (1994) Movement of respiratory CO2 in stems of loblolly pine (Pinus taeda L.) seedlings. Tree Physiology 14, 481–495. Meidner M. & Mansfield T.A. (1968) Physiology of Stomata. McGraw-Hill, New York, USA. Müller R., Baier M. & Kaiser W.M. (1991) Differential stimulation of PEP-carboxylation in guard cells and mesophyll cells by ammonium and fusicoccin. Journal of Experimental Botany 42, 215–220. Negisi K. (1974) Respiration rates in relation to diameter and age in stem or branch sections of young Pinus densiflora tree. Journal of the Japanese Forest Society 60, 380–382. Pfanz H. & Dietz K.-J. (1987) A fluorescence method for the determination of the apoplastic proton concentration in intact leaf tissue. Journal of Plant Physiology 129, 41–48. Pfanz H., Aschan G., Langenfeld-Heyser R., Wittmann C. & Loose M. (2002) Ecology and ecophysiology of tree stems: corticular and wood photosynthesis. Naturwissenschaften 89, 147– 162. Pilarski J. (1993) Intensity of oxygen production in the process of photosynthesis in shoots and leaves of lilac (Syringa vulgaris L.). Acta Physiologia Plantarum 15, 249–256. Pinelli P. & Loreto F. (2003) 12CO2 emission from different metabolic pathways measured in illuminated and darkened C3 and C4 leaves at low, atmospheric, and elevated CO2 concentration. Journal of Experimental Botany 54, 1761–1769. Rakonczay Z. (1997) Characterizing the respiration of stems and roots of three hardwood tree species in the Great Smoky Mountains. Dissertation, PhD thesis, Virginia Polytechnic Institute and State University, USA. Schaedle M. (1975) Tree photosynthesis. Annual Review of Plant Physiology 26, 101–115. Schaedle M. & Brayman A.A. (1986) Ribulose-1,5-bisphosphate carboxylase activity of Populus tremuloides Michx. bark tissues. Tree Physiology 1, 53–56. Schmidt J., Batic F. & Pfanz H. (2000) Photosynthetic performance of leaves and twigs of evergreen holly (Ilex aquifolium L.). Phyton 40, 179–190. Schönherr J. & Ziegler H. (1980) Water permeability of Betula periderm. Planta 147, 345–354. Shepard R.K. (1970) Some aspects of bark photosynthesis in bigtooth aspen (Populus grandidentata) and trembling aspen (Populus tremuloides). PhD thesis, University of Michigan, Ann Arbor, MI, USA. Sprugel D.G. & Benecke U. (1991) Measuring woody-tissue respiration and photosynthesis. In Techniques and Approaches in Forest Tree Ecophysiology (eds J.P. Lassoie & T.M. Hinckley), pp. 329–355. CRC, Boca Raton, FL, USA. Teskey R.O. & McGuire M.A. (2002) Carbon dioxide transport in xylem causes errors in estimation of rates of respiration in stems and branches of trees. Plant, Cell and Environment 25, 1571– 1577. Teskey R.O. & McGuire M.A. (2005) CO2 transported in xylem sap affects CO2 efflux from Liquidambar styraciflua and Plata-
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
1158 C. Wittmann et al. nus occidentalis stems, and contributes to observed wound respiration phenomena. Trees 19, 357–362. Von Willert D.J., Matyssek R. & Herppich W. (1995) Experimentelle Pflanzenökologie. Thieme, Stuttgart, Germany. Walker D. (1987) The Use of the Oxygen Electrode and Fluorescence Probes in Simple Measurements of Photosynthesis. Robert Hill Institute, University of Sheffield, Oxygraphics Limited, Sheffield, UK.
Wittmann C., Aschan G. & Pfanz H. (2001) Leaf and twig photosynthesis of young beech (Fagus sylvatica) and aspen (Populus tremula) trees grown under different light intensity regimes. Basic and Applied Ecology 2, 145–154. Received 31 August 2005; received in revised form 16 December 2005; accepted for publication 19 December 2005
© 2006 The Authors Journal compilation © 2006 Blackwell Publishing Ltd, Plant, Cell and Environment, 29, 1149–1158
