JXB Advance Access published online on March 20, 2007
Journal of Experimental Botany, doi:10.1093/jxb/erm038
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
RESEARCH PAPER |
Simultaneous growth and emission measurements demonstrate an interactive control of methanol release by leaf expansion and stomata
1Institute of Molecular and Cell Biology, University of Tartu, Riia 23, 51010 Tartu, Estonia
2Institute for Chemistry and Dynamics of the Geosphere:Phytosphere (ICG III), Research Centre Jülich, 52425 Jülich, Germany
3Institute of Agricultural and Environmental Sciences, Estonian University of Life Sciences, Kreutzwaldi 64, Tartu 51014, Estonia
* To whom correspondence should be addressed. E-mail: shueve{at}gmx.de
Received 6 November 2006; Revised 25 January 2007 Accepted 8 February 2007
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Emission from plants is a major source of atmospheric methanol. Growing tissues contribute most to plant-generated methanol in the atmosphere, but there is still controversy over biological and physico-chemical controls of methanol emission. Methanol as a water-soluble compound is thought to be strongly controlled by gas-phase diffusion (stomatal conductance), but growth rate can follow a different diurnal rhythm from that of stomatal conductance, and the extent to which the emission control is shared between diffusion and growth is unclear. Growth and methanol emissions from Gossypium hirsutum, Populus deltoides, and Fagus sylvatica were measured simultaneously. Methanol emission from growing leaves was several-fold higher than that from adult leaves. A pronounced diurnal rhythm of methanol emission was observed; however, this diurnal rhythm was not predominantly determined by the diurnal rhythm of leaf growth. Large methanol emission peaks in the morning when the stomata opened were observed in all species and were explained by release of methanol that had accumulated in the intercellular air space and leaf liquid pool at night in leaves with closed stomata. Cumulative daily methanol emissions were strongly correlated with the total daily leaf growth, but the diurnal rhythm of methanol emission was modified by growth rate and stomatal conductance in a complex manner. While in G. hirsutum and in F. sylvatica maxima in methanol emission and growth coincided, maximum growth rates of P. deltoides were observed at night, while maximum methanol emissions occurred in the morning. This interspecific variation was explained by differences in the share of emission control by growth processes, by stomatal conductance, and methanol solubilization in tissue water.
Key words: Fagus sylvatica, Gossypium hirsutum, leaf growth, methanol emission, Populus deltoides, stomatal conductance
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Methanol is one of the most abundant volatile organic compounds (VOCs) in the lower atmosphere, with concentrations ranging up to several tens of parts per billion (ppb; Riemer et al., 1998; Singh et al., 2000; Schade and Goldstein, 2001; Jacob et al., 2005). Since methanol has a major impact on OH radical concentrations as well as on photochemical ozone formation, efforts have been made to characterize methanol sources and emission rates. Early field and laboratory measurements provided evidence of a dominant role for biogenic processes in methanol release. Fehsenfeld et al. (1992) found methanol as the major VOC in a pine forest, and MacDonald and Fall (1993) reported methanol emissions from leaves. Nemecek-Marshall et al. (1995) showed that growing leaves emit much higher amounts of methanol than adult leaves, and suggested that growth processes play an important role in these emissions. Galbally and Kirstine (2002) further estimated the source strength for methanol emissions originating from growth processes to be 100–128 Tg year–1, i.e. about two-thirds of total global methanol emissions.
Growth processes are associated with structural changes of cell walls and demethylation of pectin, a likely source of leaf methanol (Fall and Benson, 1996). Accordingly, methanol emissions have been observed during different processes that include changes in the cell wall structure, such as expansion growth of roots, leaves, and fruits (Fall and Benson, 1996), growth and maturation of seeds (Obendorf et al., 1990), destruction of cell walls and generation of intercellular air spaces (Nemecek-Marshall et al., 1995; Fall and Benson, 1996), and during leaf abscission and ageing of plant tissues (Harriman et al., 1991). Although the dominant role of growth processes in methanol production is known, information on the mechanisms of methanol release from plants is scarce (Galbally and Kirstine, 2002, and references therein). Leaf growth varies greatly during the day and has a pronounced day/night cycle, and the exact timing of these variations differs among plant species (Walter and Schurr, 2005). Therefore, it is important to determine how the dynamics of leaf growth is reflected in methanol emission.
There is further evidence that gas-phase diffusion conductance (GG), i.e. stomatal conductance, can significantly modify the methanol emission rate (MacDonald and Fall, 1993; Nemecek-Marshall et al., 1995; Niinemets and Reichstein, 2003b). In several volatile plant compounds such as isoprene, the emission flux [
=GG(Pi–Pa)/P, where Pi is the compound internal pressure, Pa the atmospheric partial pressure, and P is total air pressure] is not controlled by GG (Fall and Monson, 1992). This is because after a reduction in GG, Pi–Pa rises essentially immediately, compensating for the decrease in conductance (Fall and Monson, 1992; Niinemets et al., 2004). Differently from isoprene, methanol has a high water solubility and, therefore, a Pi–Pa rise requires a large time-consuming change in liquid-phase methanol concentration, and accordingly methanol emission can be temporarily curbed by low gas-phase diffusion conductance (Niinemets and Reichstein, 2003b).
Hence, the dynamics of methanol emission from plants should depend on growth processes as well as on stomatal conductance. So far, simultaneous measurements of the diurnal variation of growth, stomatal conductance, and methanol emission are lacking. In this study, detailed growth measurements during the entire day were combined with simultaneous measurements of methanol emission and stomatal conductance to determine the extent to which the emission control is shared between growth and diffusion processes. These measurements were conducted in three species that have different variations in leaf growth activities during a day/night cycle. In cotton (Gossypium hirsutum L.), the highest leaf growth rates are found in the morning, while in poplar (Populus deltoides Bartr. ex Marsh) and beech (Fagus sylvatica L.) leaf growth mainly occurs at night. In all three species, stomata are open during the day and closed at night, but the efficiency of stomatal control at night varies among the species. The working hypothesis was that in growing leaves, the diurnal variation in methanol emission is mainly controlled by the leaf expansion rate and that diurnal changes in stomatal conductance can result in phase shifts between growth rate and methanol emission rate.
|
Materials and methods |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Plant material
Seeds of cotton plants (G. hirsutum L. var. Stoneville) were germinated in a commercial mixture of soil, peat, and compost (Pikiererde, Plantaflor, Vechta, Germany). After germination, the plants were cultivated individually in standard soil (Einheitserde, type ED 73) at a temperature of 28 °C (day)/25 °C (night), with 60–85% relative humidity, 14 h photoperiod, and a PAR of 300 µmol m–2 s–1. Gossypium hirsutum plants were used in the experiments when they were between 9 and 12 weeks old.
Poplar (P. deltoides Bartr. ex. Marsh, clone S7c8) plants propagated from cuttings were grown in 1.0 l pots in a glasshouse and watered daily. The photoperiod in the glasshouse was maintained at 16 h day/8 h night. Illumination in the glasshouse (SON-T AGRO 400, Philips) was automatically turned on when the ambient light intensity outside the glasshouse was <400 µmol m–2 s–1 during daytime. The temperature in the glasshouse was 25±1 °C during the day and 18±1 °C during the night.
Two- to three-year-old seedlings of beech (F. sylvatica L.) were grown outdoors in unshaded conditions. The plants passed through a normal winter dormancy cycle outdoors and were transferred indoors for the experiments in spring. Leafless seedlings were transferred to the gas-exchange measurement system, and the bud break occurred in the gas-exchange system during the experiments.
Measurement of methanol emissions, water vapour, and CO2 exchange
The experiments were conducted in continuously stirred tank reactors (CSTRs) made of glass. The aboveground parts of whole plants were enclosed in the chamber, and chambers of different sizes were used for the experiments, depending on plant size. For experiments involving larger plants, chambers with volumes of 164 l and 1100 l were used, while a 5 l chamber was employed for experiments with smaller plants (see Beauchamp et al., 2005 for further details of the chambers used; Heiden et al., 1999, 2003; Wildt et al., 2003).
Except where noted, day length was 12 h in the experiments with G. hirsutum and 11 h in the experiments with P. deltoides. In F. sylvatica that was transferred from outdoor conditions, there was 1 h illumination at 
50 µmol m–2 s–1 in the morning and in the evening simulating twilight, and an 11 h day illumination period in between the ‘twilight’ periods. Air temperature was maintained at 28 °C (day)/25 °C (night). Leaves were well coupled to ambient atmosphere such that leaf temperatures deviated from ambient air temperature at most by ±2 °C. During the day, the photosynthetic quantum flux density was 800±50 µmol m–2 s–1 in the 164 l chamber, 480±20 µmol m–2 s–1 in the 1100 l chamber, and 200±10 µmol m–2 s–1 in the 5 l chamber. Dew point in the chambers was between 15 °C and 20 °C, depending on plant transpiration.
Ambient air was purified by a palladium catalytic converter and an adsorptive drying device (KEA 70, Ecosorb filter, Zander, Essen, Germany). The air was then pumped through the chambers, with Teflon fans providing homogeneous mixing of the chamber air. The flow rates were between 1.2 mmol s–1 and 29.8 mmol s–1, depending on the size of the plants and the chamber used for the experiments. These flow rates resulted in mean residence times of the air in the chamber of between 1 min and 5 min in most cases and up to 28 min in the largest F. sylvatica plants in the 1100 l chamber. Selection of appropriate dew point and flux rates assured that no condensation of water occurred in the chamber and in the tubing.
Absolute mixing ratios of CO2 and H2O at the chamber inlet were measured with infrared gas analysers (model URAS 3G, Hartmann & Braun, Frankfurt a. M., Germany) and dew-point mirrors (MTS MK1, Walz, Effeltrich, Germany), while additional infrared gas analysers (BINOS 100, Rosemount Analytical, Hasselroth, Germany) were used to measure the difference in CO2 and H2O mixing ratios between the chamber inlet and outlet (Schuh et al., 1997; Heiden et al., 1999).
VOC concentrations were determined using an on-line gas chromatograph (HP 6890, Hewlett Packard) with a quadrupole mass selective detector (HP-MSD 5973, Hewlett Packard) (Folkers, 2002). The system is optimized to detect quantitatively various volatiles ranging from oxygenated compounds such as methanol, acetone, C6 aldehydes, and alcohols to hydrophobic compounds such as isoprene and monoterpenes. For a full spectrum of compounds, the time resolution was 45 min. In the experiments described here, mainly methanol emissions, and in some cases emissions of other low-molecular oxygenated compounds such as acetone, were measured, and the time resolution achieved was 25 min.
The system was calibrated as described previously (Folkers, 2002; Heiden et al., 2003). A small vial closed by a Teflon membrane was filled with the specific compound. This vial was stored in a temperature-controlled glass vessel flushed with pure nitrogen. VOC diffusing through the membrane from the vial into the vessel was diluted in the nitrogen stream and then carried out of the glass vessel by the nitrogen stream. To obtain concentrations in the ppb range, the nitrogen flow was diluted in air by a dynamic system. The concentration of the specific volatile compound released from the calibration source was determined from the mass loss rates and the dilution fluxes. The overall accuracy of the emission measurements of methanol and other oxygenated compounds was estimated to be better than 30%, and the reproducibility of the measurements was between 5% and 10%. When operated in the mode with high temporal resolution, the detection limit for methanol was about 500 ppt.
Calculation of transpiration and assimilation rates, and emission fluxes
Leaf transpiration, photosynthesis rate, and stomatal conductance were calculated according to von Caemmerer and Farquhar (1981). The emission rate of a compound X (X) was calculated using the measured mixing ratios of a compound X at the chamber outlet and inlet as:
 | (1) |
In G. hirsutum and P. deltoides, either a few expanding leaves were present during the 4–8 d the plant was enclosed in the chamber, or leaf area growth was measured continuously during the emission measurements, making it possible to determine the appropriate total leaf area in all cases. In F. sylvatica, the emissions were measured for 14–21 d. When the plants were enclosed in the chamber, the bud break had not yet occurred and, after bud break, leaf growth occurred at many points in the seedling canopy simultaneously. Due to uncertainties with total leaf area at various times during the experiment in F. sylvatica, differences in VOC and H2O mixing ratios between chamber inlet and outlet are reported for this species.
Measurements of leaf growth
For simultaneous leaf growth and methanol emission measurements in G. hirsutum and P. deltoides, a young apical leaf was selected in each case. The plant was then enclosed in the chamber, and all other leaves remaining in the chamber were removed. The rate of leaf expansion growth (expressed as the change in leaf area in % h–1) and emission of VOCs were then measured continuously for several days.
A digital image sequence processing method was employed for quantification of expansion growth (Schmundt et al., 1998; Walter et al., 2002a, b; Matsubara et al., 2006). Leaves were fixed gently to the focal plane of a CMOS camera (Flea-BW, Point Grey Research, Vancouver, Canada) and were illuminated with infrared diodes (880 nm) to ensure constant image brightness throughout night and day. Near-infrared images of the leaf were captured every 120 s on the adaxial side of the leaf through an interference filter (880 nm, Schott, Mainz, Germany).
Image sequences were evaluated with algorithms based on a structure–tensor approach (Schmundt et al., 1998). According to these algorithms, the movement velocities of all structures consistently moving within an image sequence of a growing leaf are calculated. Within the greyscale image sequence, areas of the foliage with characteristic grey values are shifted in time because of the expansion of the leaves. To calculate the distribution of relative growth rate (RGR) within the foliage, displacement vector fields for each image are calculated. Any leaf location with a suitable grey value contrast to its neighbourhood results in oriented grey value structures in image stacks. The arrangements of these trajectories relative to the spatio-temporal neighbourhood of a central pixel are analysed via an optical flow approach. This analysis provides a velocity vector for the motion of the central pixel (Walter et al., 2002a, b). RGRs were obtained by tracking time-dependent deformation of a polygonal area of interest selected within the image. Twenty-five minute averages of RGR were calculated to synchronize the growth data with the sampling interval of the gas chromatograph. In general, active growth alternates with periods of arrested growth or even periods with negative growth, during which the leaf tissues are shrinking (Lai et al., 2005). In this study, spikes of negative growth were occasionally visible in G. hirsutum leaves at night.
The growth of F. sylvatica leaves was monitored throughout 6–8 d immediately after bud break using linear voltage displacement transducers (LVDTs; Althen, Kelkheim, Germany) that were connected to the leaf tips via nylon threads and recorded the increase in leaf length electronically. Growth of four leaves from a single plant was recorded under the same climatic conditions as those under which the methanol emission measurements were conducted. Average values for leaf length increase per hour were calculated from these data.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Diurnal rhythm of gas exchange and methanol emission in G. hirsutum
Variations in gas-exchange rates and methanol emission during the day were studied in 13 different experiments with a variable number of G. hirsutum plants (1–4 plants in the individual experiments) and qualitatively similar patterns were observed in all cases. After switching on the light, leaf transpiration rates rapidly increased and stayed at a high level during most of the day, with a minor depression in the evening (Fig. 1). Under the experimental conditions chosen here, with constant temperature during illumination and high air flow rates, time courses of transpiration and stomatal conductance were generally very similar. Diurnal variation in net assimilation rates followed the same time course as transpiration. Assimilation rates during the daytime usually varied between 5 µmol m–2 s–1 and 8 µmol m–2 s–1, with an average of 6.8 µmol m–2 s–1 during the day shown in Fig. 1.
|
Emissions of acetone and most other oxygenated VOCs essentially followed the leaf transpiration rate (data not shown). Diurnal variation of methanol emission differed substantially from the simple time course observed in other compounds. During the first hour after the onset of dark phase, methanol emissions were low, but significantly above the detection limit (Fig. 1). After a few hours in darkness, the emissions increased according to biphasic kinetics. The first phase, lasting
5 h (4–9 h in darkness), was characterized by a slow increase in the methanol emission rate, while the emissions increased more rapidly between 9 h and 12 h in darkness. During this phase, stomata opened slightly in the dark. A sharp peak of emission in the morning coincided with the light-dependent increase in stomatal conductance and transpiration (Fig. 1). Subsequently, methanol emissions decreased nearly exponentially with decay times between 2 h and 10 h, while the transpiration rate was essentially constant. When the light was switched off at the end of day, a detectable drop of methanol emission was observed (Fig. 1). To gain further insight into the influences of light on methanol emissions, different daily light regimes were employed. During continuous illumination, there was still a pronounced diurnal rhythm in methanol emission (Fig. 2a). However, the conspicuous morning peak of methanol emission that accompanied the onset of the period of illumination in the morning was absent during continuous illumination (cf. Figs 2a and 1). Diurnal rhythm in stomatal conductance can be inferred from the pattern of transpiration, showing a weaker stomatal response in days with continuous illumination than during a standard day/night light cycle (Fig. 2a). When the permanent illumination was changed back to the normal day/night illumination rhythm (12 h light/12 h darkness), the emission peak of methanol after onset of light was visible again (Fig. 2a). Despite the different kinetic patterns between days with permanent illumination and with day/night illumination rhythm, the total amount of methanol emitted over 24 h was not different between the two light regimes (Fig. 2b).
|
Rapid changes in the environment induced stomatal oscillations in G. hirsutum, as is common in many plants (Farquhar and Cowan, 1974; Rose and Rose, 1994). A sample response of transpiration rate and methanol emission rate after abrupt darkening during the light period followed by illumination is shown in Fig. 3. This rapid disturbance induced oscillations in stomatal aperture with a period of about 30 min (Fig. 3), which were followed by oscillations of methanol emission.
|
Methanol emission in relation to leaf age and growth in G. hirsutum
The experiments described suggested that stomata exert a major control over foliar methanol emissions. However, the divergence of emission and transpiration kinetics during the day indicated that other factors also importantly modify methanol emissions. Comparison of methanol emissions from young growing leaves and from older adult leaves indicated several-fold larger emissions from growing leaves (Fig. 4). While the emissions from the plants with fully expanded leaves were lower, the diurnal time courses of the emissions in young and old leaves were similar, with a pronounced morning peak concomitant with increases in stomatal aperture. Reduction of methanol emission after removal of growing leaves implied that
75% of the methanol emission was due to leaf growth (Fig. 4).
|
Simultaneous measurements of expansion growth and methanol emission demonstrated that the diurnal courses of these two processes were broadly overlapping (Fig. 5). To discriminate between different factors producing this overlap (simultaneous light-dependent triggering of growth and stomatal opening, endogenous rhythm), the duration of illumination was reduced from 12 h to 6 h at day 3 (Fig. 5), after which standard 12 h light/12 h dark rhythm was restored, but with a phase shift of 6 h. The phase shift in light had little effect on the diurnal course of expansion growth rate (Fig. 5). The methanol emission also remained coupled to growth rate. However, on the day immediately after phase shift, the conspicuous peak in methanol emission directly after the onset of illumination was absent. In fact, the stomata opened on that day with a slower rate due to a pronounced memory effect (endogenous rhythm; data not shown, see Fig. 2a).
|
Methanol emission in relation to leaf growth in P. deltoides and F. sylvatica
In P. deltoides, the expansion growth had a maximum in the early hours of darkness after the illumination period (Fig. 6), and there was a time shift of about 12 h between the maxima in growth and methanol emission (Fig. 6). Despite different diurnal growth kinetics, the diurnal courses of methanol emission in P. deltoides were similar to those in G. hirsutum. In particular, the peak appearing together with the increase of stomatal conductance in the morning when the lamps were turned on was observed (Fig. 6).
|
The maximum emission rate during the peaks and the steady-state emission rate decayed from day to day in P. deltoides—a feature comparable with the time-dependent decline in leaf growth rate in this species. Although the expansion growth and methanol emission exhibited out-of-phase kinetics, the cumulative daily methanol emissions correlated closely with total daily leaf growth (Fig. 7).
|
In F. sylvatica, bud break occurred on day 3 after enclosure of trees in the chamber. After bud break, measurable methanol emissions were observed (Fig. 8a). In contrast to the other species, foliage of F. sylvatica emitted methanol predominantly during the dark phase (Fig. 8a). An emission peak concomitant with the onset of illumination became apparent on day 7. Growth measurements indicated that leaf growth was almost entirely restricted to the dark phase in this species (Fig. 8b).
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Origin of methanol emitted from plants
Plant emissions are believed to constitute the largest source of atmospheric methanol (Singh et al., 2000). The average emission rate in 11 species studied by MacDonald and Fall (1993) varied between 0.005 and 0.15 nmol (g dry mass)–1 s–1. In G. hirsutum, the 24 h average methanol emission rate (e.g. Fig. 2b) in the same units was 0.011–0.1 nmol (g dry mass)–1 s–1. However, the peak emissions were as high as 0.4 nmol (g dry mass)–1 s–1 (Fig. 1). These average and maximum methanol emission rates observed in this study are comparable with those found in other studies (Nemecek-Marshall et al., 1995; Warneke et al., 2002; Karl et al., 2003).
Galbally and Kirstine (2002), using the data from MacDonald and Fall (1993), estimated that the plant carbon loss due to methanol emission is of the order of 0.11–0.16% of total carbon fixed in photosynthesis. In the experiments shown here, a relatively high percentage of CO2 fixed in photosynthesis was emitted as methanol. This percentage (daily average) varied between 0.05% and 0.55% in G. hirsutum, 0.04% and 0.3% in P. deltoides, and 0.03% and 0.2% in F. sylvatica. Prolonged or shortened CO2 uptake by changes of the day–night phase did not noticeably influence the amount of methanol emitted (e.g. Figs 2 and 5), demonstrating that methanol emissions are not directly associated with daily leaf net CO2 uptake. Similar observations were made by Nemecek-Marshall et al. (1995), where even CO2-free air did not eliminate methanol emissions.
Experiments with old versus young leaves (Fig. 4) indicated that 75% of the methanol emission could be ascribed to leaf growth. Growth and the demethylation of pectin has long been proposed as a source of methanol emission (Nemecek-Marshall et al., 1995). During growth processes, chemical bonds between cell wall macromolecules such as celluloses and pectins have to be cleaved and cross-linked again, when new cell wall material is incorporated into the extending apoplast (Cosgrove, 1999). Other processes associated with changes in cell wall structure, such as seed maturation and fruit ripening, have also been accompanied by methanol emissions (Obendorf et al., 1990; Frenkel et al., 1998; Koch et al., 1999; Lee et al., 2001). The demethylation of methyl esters of polygalacturonans in cell wall pectin is catalysed by pectin methylesterases (PMEs) that are ubiquitous cell wall enzymes. In dicots, these enzymes are involved in important developmental processes including cellular adhesion and stem elongation (Micheli, 2001; Willats et al., 2001; Tiznado-Hernández et al., 2004). Other significant sources of plant-generated methanol include protein repair pathways (Fall and Benson, 1996), or stress-induced emissions that probably come from cell wall degradation (Fukui and Doskey, 1998; Warneke et al., 2002; von Dahl et al., 2006).
Methanol emission is partially stomata dependent and has a pronounced peak in the early morning
Earlier work has suggested that foliar methanol emission occurs through stomata and is controlled by stomatal conductance (MacDonald and Fall, 1993; Nemecek-Marshall et al., 1995), but leaf growth rate was not measured in these studies. As growth may vary simultaneously with transpiration rate as in G. hirsutum in this study, measurements of emission and transpiration rates alone are not always conclusive. Even artificial reduction in stomatal conductance by the plant hormone abscisic acid (ABA) may not be convincing (Nemecek-Marshall et al., 1995) as ABA can also affect expansion growth. The methanol emission rate from the plant leaves can be described as in =GG(Pi–Pa)/P, with F/A being replaced by GG, the stomatal conductance for methanol (Niinemets et al., 2002; Niinemets and Reichstein, 2003a, b). When GG decreases, and the rate of methanol synthesis remains constant, an increase of Pi–Pa may compensate for reduction in stomatal conductance, and there would be no stomatal control in a steady state. However, the key issue with stomatal control of VOC emissions is how fast is the rise in the partial pressure difference. For water-soluble compounds such as methanol, Pi–Pa rises relatively slowly, and the stomata may control emissions in a non-steady state. The diffusion flux from the site of methanol synthesis in the cell walls to the internal air space is given as:
 | (2) |
=m. However, for compounds with low H (large solubility), a large change in liquid-phase concentration is needed for a certain change in Pi and, accordingly, stomata may temporarily control the emission rate. Henry's law constant for methanol is 0.46 Pa m3 mol–1 at 25 °C, implying that methanol is highly soluble in water (for comparison, the Henry's law constant for the hydrophobic compound isoprene is 7780 Pa m3 mol–1 at 25 °C; see Niinemets and Reichstein, 2003b). Three pieces of evidence suggested stomatal control of methanol emissions. In the morning, changes in transpiration were associated with significant changes in methanol emission (Fig. 1), although these diurnal changes were over-ridden by growth dynamics (Figs 1 and 5 versus Figs 6 and 8). Secondly, the oscillations in stomatal conductance were accompanied by concomitant oscillations in methanol emission (Fig. 3).
A burst of methanol emission after switching on the light provided the third piece of evidence for the partial control of methanol emissions by stomatal conductance. In cotton, the peak of leaf growth tended to coincide with stomatal opening, probably increasing the morning peak of methanol emission (Figs 1, 2, and 5). Nevertheless, the data obtained with P. deltoides, exhibiting peaks of methanol emission that were completely unrelated to growth (Fig. 6), clearly demonstrated that the methanol emission bursts were not only associated with growth. During periods of low stomatal conductance, little methanol is emitted and a large liquid-phase pool of methanol can be built up within the leaf (Niinemets and Reichstein, 2003b). In the morning, when the stomata open, accumulated methanol is emitted in a burst (Niinemets et al., 2002, 2004; Niinemets and Reichstein, 2003b; for a review of these model studies, see Fall, 2003). In a very similar mechanism, internal methanol pools are released in a short and sharp peak after wounding (Loreto et al., 2006). Morning bursts of emission were observed in all species investigated here and were also reported by Nemecek-Marshall et al. (1995). In flux measurements in Medicago sativa meadows, similar morning bursts were also observed (Warneke et al., 2002). Although the authors attributed these morning bursts of methanol emissions to accumulation of methanol in dew and release from dew during dew evaporation, these bursts may also result from stomatal opening.
Key issues with the stomata-dependent morning burst are how large is the pool and how large is the change in stomatal conductance. In Phaseolus vulgaris, Nemecek-Marshall et al. (1995) measured free methanol pools between 310 nmol g–1 fresh mass in the oldest leaves, and 840 nmol g–1 fresh mass in young, fully expanded leaves, while the steady-state emissions rates for the same leaves were 0.52 nmol (g dry mass)–1 s–1 for the youngest leaves and 0.09 nmol (g dry mass)–1 s–1 for old leaves, indicating that the pools can be large enough to sustain a burst-like methanol emission without a need for de novo synthesis during the burst. In this study, the emission slowly increased throughout the night when stomata were closed, demonstrating that methanol did accumulate in the liquid phase. Methanol accumulation could be at least partly responsible for the burst (Fig. 1). In several manipulative experiments, morning bursts were absent. This happened in days when the plants were continuously illuminated and the change in stomatal conductance was less pronounced (Fig. 2a) as well as in the day immediately following the reduction of day-length, when the amount of methanol accumulated was less and stomatal response in the morning was delayed (Fig. 5).
An interesting phenomenon observed in this study was the switch-like reduction of the methanol emission rate after darkening the plant at the end of the light period (e.g. Figs 1 and 4). This reduction in emission rate was much faster than reduction in stomatal conductance. However, concomitantly with light, temperature was also reduced, and it is suggested that this reduction in emission reflected enhanced gas-phase diffusion limitation because of an increased methanol solubility in cell wall water (H changes from 0.55 Pa m3 mol–1 at 28 °C to 0.46 Pa m3 mol–1 at 25 °C). The opposite change in solubility in the morning may have moderately contributed to the emission burst after switching on the light.
Control of methanol emissions by leaf age and growth
Methanol emission drastically declined in old leaves, confirming the previous studies (Nemecek-Marshall et al., 1995). Despite a clear age-dependent decline, methanol emission was actually maximal from leaves of the fourth node rather than from the youngest leaves of P. vulgaris (Nemecek-Marshall et al., 1995). In this study, methanol emission declined in a log-linear fashion with increasing leaf age in P. deltoides (Fig. 6). However, the plants were measured after bud burst in this experiment when the leaves had attained a size of 4 cm2. In F. sylvatica, in which the bud burst occurred in the chamber, methanol emissions were maximized at day 7, comparable with the study of Nemecek-Marshall et al. (1995). Given that stomata develop and mature simultaneously with leaf expansion growth (Willmer, 1983), delayed methanol emissions from very young leaves can be associated with the lack of functional stomata.
To characterize the relationship between methanol emission and leaf growth quantitatively, additional experiments with simultaneous measurements of leaf growth and methanol emissions were conducted with plants of G. hirsutum, P. deltoides, and F. sylvatica. In these species, the maximum growth activity occurred at different times of the day.
Leaf growth rate undergoes strong diurnal changes that are associated with differential timing of enhanced turgor and increasing availability of carbohydrates from photosynthesis (daytime growth maximum) or starch breakdown (night/early morning maximum) (Walter et al., 2002a; Walter and Schurr, 2005). Expansion rates of, for example, Ricinus leaves have a distinct maximum during the second half of the dark period and in the early morning; a timing that is comparable with that of G. hirsutum leaves (Walter et al., 2002a; Walter and Schurr, 2005). As noted previously (Walter et al., 2005; Matsubara et al., 2006), the maximum growth rate was observed in the early hours of the night in P. deltoides, and throughout the entire night in F. sylvatica, when the stomata were essentially closed.
Species variation in growth timing resulted in major differences in the timing of methanol emission and association of the emissions with RGR. In G. hirsutum, the peaks of methanol emission and growth rate were almost synchronized. As discussed above, the morning peak can be partly associated with rapid modifications in stomatal conductance and release of methanol accumulated at night. However, experiments involving continuous illumination (Fig. 2a) demonstrated that the emission maximum, albeit less pronounced, is also present when changes in stomatal conductance are less. Direct growth measurements (Fig. 5) further suggested that the morning peak and pronounced daily dynamics of methanol emissions are significantly driven by the growth-dependent variability in methanol production rate. Thus, two independent processes interacted to cause the morning emission peak in G. hirsutum. The growth maximum in the early morning resulted in an emission maximum that was further superimposed by the emission peak caused by opening of stomata and the release of methanol dissolved in water.
In P. deltoides, there was a shift of about 12 h between the maxima of growth rate and methanol emission (Fig. 6), while the cumulative daily methanol emissions correlated well with daily growth (Fig. 7). Methanol produced during the phase of maximal growth activity in the beginning of the night may be dissolved in tissue water and is released upon opening the stomata the next morning, explaining the morning burst of emission, i.e. the methanol source would be essentially the same as in cotton, but with delayed emissions due to stomatal limitations and intermediate storage in water. An alternative explanation for the discrepancy between emission and growth dynamics is that leaf expansion and chemical modification of pectins are temporally separated in this species. Plant growth consists of a series of events involving cell wall relaxation, cell expansion, cell wall thickening, and hardening (Cosgrove, 2000). Pectin demethylation by pectin methylesterases leading to formation of Ca2+ cross-links occurs during cell wall hardening (Cosgrove, 2005), and must not necessarily occur simultaneously with the bulk of leaf expansion. Nevertheless, the baseline (minimum) methanol emissions in this species were somewhat larger than in G. hirsutum and F. sylvatica (cf. Fig. 6 and Figs 5 and 8), and also correlated with leaf growth rate on the specific day, suggesting that the control over methanol emission may be shared between expansion growth and stomata, similarly to what occurs in G. hirsutum.
Differently from the two other species, the leaves of F. sylvatica emitted the bulk of methanol at night when most leaf expansion growth occurred, despite the fact that stomatal conductance was low at night (Fig. 8). At the same time, the stomatal control was relatively weak in these young leaves, as the result of which the characteristic morning peak was absent until day 7, and the day/night cycle of emissions was mainly driven by the expansion growth rate.
Conclusions
Taken together, the data suggest that growth processes exert a major control over methanol release, but also that the correlations between the expansion growth rate and methanol emission are modified by stomata, and possibly also by the time lags in the process sequence from cell wall expansion growth to cell wall hardening. Methanol, probably originating from pectin demethylation, is released into cell wall water and leaf intercellular air space. The rate of emission into the atmosphere depends on stomatal conductance and the rate of methanol production in growth processes. In conditions of closed stomata, water-soluble methanol accumulates in cell wall water and can be released in a burst-like fashion after rapid stomatal opening. It is suggested that the complex daily dynamics of methanol emission can be explained by the diurnal kinetics of growth and the share of control of emission between stomatal and growth processes.
|
Acknowledgements |
|---|
The study was supported by European Science Foundation project VOCBAS (Volatile organic compounds in biosphere–atmosphere system), the Estonian Science Foundation (Grant 5702), the Estonian Ministry of Education and Science (grant 0182468As03), and the Estonian Academy of Sciences.
|
References |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Beauchamp J, Wisthaler A, Hansel A, Kleist E, Miebach M, Niinemets Ü, Schurr U, Wildt J. (2005) Ozone induced emissions of biogenic VOC from tobacco: relations between ozone uptake and emission of LOX products. Plant, Cell and Environment 28 1334–1343.
Cosgrove DJ. (1999) Enzymes and other agents that enhance cell wall extensibility. Annual Review of Plant Physiology and Plant Molecular Biology 50 391–417.[CrossRef][Web of Science][Medline]
Cosgrove DJ. (2000) Expansive growth of plant cell walls. Plant Physiology and Biochemistry 38 109–124.[CrossRef][Web of Science][Medline]
Cosgrove DJ. (2005) Growth of the plant cell wall. Nature Reviews Molecular Cell Biology 6 850–861.[CrossRef][Web of Science][Medline]
Fall R. (2003) Abundant oxygenates in the atmosphere: a biochemical perspective. Chemical Reviews 103 4941–4952.[CrossRef][Web of Science][Medline]
Fall R and Benson AA. (1996) Leaf methanol—the simplest natural product from plants. Trends in Plant Science 1 296–301.
Fall R and Monson RK. (1992) Isoprene emission rate and intercellular isoprene concentration as influenced by stomatal distribution and conductance. Plant Physiology 100 987–992.
Farquhar GD and Cowan IR. (1974) Oscillations in stomatal conductance. Plant Physiology 54 769–772.
Fehsenfeld F, Calvert J, Fall R, et al. (1992) Emissions of volatile organic compounds from vegetation and the implications for atmospheric chemistry. Global Biogeochemical Cycles 6 389–430.
Folkers A. (2002) Sauerstoffhaltige flüchtige organische Verbindungen in der Troposphäre: Entwicklung und Anwendung einer gaschromatographischen Nachweismethode. Berichte des Forschungszentrums Jülich. Jül-3998Jülich Forschungszentrum Jülich.
Frenkel C, Peters JS, Tieman DM, Tiznado ME, Handa AK. (1998) Pectin methylesterase regulates methanol and ethanol accumulation in ripening tomato (Lycopersicon esculentum) fruit. Journal of Biological Chemistry 273 4293–4295.
Fukui Y and Doskey PV. (1998) Air–surface exchange of nonmethane organic compounds at a grassland site: seasonal variations and stressed emissions. Journal of Geophysical Research–Atmospheres 103 13153–13168.
Galbally IE and Kirstine W. (2002) The production of methanol by flowering plants and the global cycle of methanol. Journal of Atmospheric Chemistry 43 195–229.
Harriman RW, Tieman DM, Handa AK. (1991) Molecular cloning of tomato pectin methylesterase gene and its expression in rutgers, ripening inhibitor, non-ripening, and never ripe tomato fruits. Plant Physiology 97 80–87.
Heiden AC, Hoffmann T, Kahl J, et al. (1999) Emission of volatile organic compounds from ozone-exposed plants. Ecological Applications 9 1160–1167.
Heiden AC, Kobel K, Langebartels C, Schuh-Thomas G, Wildt J. (2003) Emissions of oxygenated volatile organic compounds from plants. Part I: emissions from lipoxygenase activity. Journal of Atmospheric Chemistry 45 143–172.
Jacob DJ, Field BD, Li Q, Blake DR, de Gouw J, Warneke C, Hansel A, Wisthaler A, Singh HB, Guenther A. (2005) Global budget of methanol: constraints from atmospheric observations. Journal of Geophysical Research–Atmospheres 110 D08303 08310.01029/02004JD005172.
Karl T, Guenther A, Spirig C, Hansel A, Fall R. (2003) Seasonal variation of biogenic VOC emissions above a mixed hardwood forest in Northern Michigan. Geophysical Research Letters 30 2186 doi:2110.1029/2003GL018432.
Koch JL, Horbowicz M, Obendorf RL. (1999) Methanol, pectin and pectin esterase changes during soybean seed maturation. Seed Science Research 9 311–320.
Lai I-L, Scharr H, Chavarria-Krauser A, Küsters R, Wu J-T, Chou C-H, Schurr U, Walter A. (2005) Leaf growth dynamics of two congener gymnosperm tree species reflect the heterogeneity of light intensities given in their natural ecological niche. Plant, Cell and Environment 28 1496–1505.
Lee P-C, Taylor AG, Zhang M, Esashi Y. (2001) Volatile compounds and accumulation of acetaldehyde–protein adducts in relation to seed quality and storage conditions. Journal of New Seeds 2 59–76.
Loreto F, Barta C, Brilli F, Nogues I. (2006) On the induction of volatile organic compound emissions by plants as consequence of wounding or fluctuations of light and temperature. Plant, Cell and Environment 29 1820–1828.[CrossRef][Medline]
MacDonald RC and Fall R. (1993) Detection of substantial emissions of methanol from plants to the atmosphere. Atmospheric Environment 27A 1709–1713.
Matsubara S, Hurry V, Druart N, Benedict C, Janzik I, Chavarría-Krauser A, Walter A, Schurr U. (2006) Nocturnal changes in leaf growth rate of Populus deltoides are controlled by cytoplasmic growth. Planta 223 1315–1328.[CrossRef][Web of Science][Medline]
Micheli F. (2001) Pectin methylesterases: cell wall enzymes with important roles in plant physiology. Trends in Plant Science 6 414–419.[CrossRef][Web of Science][Medline]
Nemecek-Marshall M, MacDonald RC, Franzen JJ, Wojciechowski CL, Fall R. (1995) Methanol emission from leaves. Enzymatic detection of gas-phase methanol and relation of methanol fluxes to stomatal conductance and leaf development. Plant Physiology 108 1359–1368.[Abstract]
Niinemets Ü, Loreto F, Reichstein M. (2004) Physiological and physico-chemical controls on foliar volatile organic compound emissions. Trends in Plant Science 9 180–186.[CrossRef][Web of Science][Medline]
Niinemets Ü and Reichstein M. (2003a) Controls on the emission of plant volatiles through stomata: a sensitivity analysis. Journal of Geophysical Research–Atmospheres 108 4211 doi:4210.1029/2002JD002626.
Niinemets Ü and Reichstein M. (2003b) Controls on the emission of plant volatiles through stomata: sensitivity or insensitivity of the emission rates to stomatal closure explained. Journal of Geophysical Research–Atmospheres 108 4208 doi:4210.1029/2002JD002620.
Niinemets Ü, Reichstein M, Staudt M, Seufert G, Tenhunen JD. (2002) Stomatal constraints may affect emission of oxygenated monoterpenoids from the foliage of Pinus pinea. Plant Physiology 130 1371–1385.
Obendorf RL, Koch JL, Gorecki RJ, AR A, Aveni MT. (1990) Methanol accumulation in maturing seeds. Journal of Experimental Botany 41 489–495.
Riemer D, Pos W, Milne P, et al. (1998) Observations of nonmethane hydrocarbons and oxygenated volatile organic compounds at a rural site in the southeastern United States. Journal of Geophysical Research–Atmospheres 103 28111–28128.
Rose MA and Rose MA. (1994) Oscillatory transpiration may complicate stomatal conductance and gas-exchange measurements. HortScience 29 693–694.
Schade GW and Goldstein AH. (2001) Fluxes of oxygenated volatile organic compounds from a ponderosa pine plantation. Journal of Geophysical Research 106 3111–3123.
Schmundt D, Stitt M, Jähne B, Schurr U. (1998) Quantitative analysis of local growth rates of dicot leaves at high temporal and spatial resolution, using image sequence analysis. The Plant Journal 16 505–514.
Schuh G, Heiden AC, Hoffmann T, Kahl J, Rockel P, Rudolph J, Wildt J. (1997) Emissions of volatile organic compounds from sunflower and beech: dependence on temperature and light intensity. Journal of Atmospheric Chemistry 27 291–318.
Singh HB, Chen Y, Tabazadeh A, et al. (2000) Distribution and fate of selected oxygenated organic species in the troposphere and lower stratosphere over the North Atlantic. Journal of Geophysical Research–Atmospheres 105 3795–3805.
Tiznado-Hernández M-E, Gaffe J, Handa AK. (2004) Isolation and study of a ubiquitously expressed tomato pectin methylesterase regulatory region. Electronic Journal of Biotechnology 7 9–29.
von Caemmerer S and Farquhar GD. (1981) Some relationships between the biochemistry of photosynthesis and the gas exchange of leaves. Planta 153 376–387.
von Dahl C, Hävecker M, Schlögl R, Baldwin IT. (2006) Caterpillar-elicited methanol emission: a new signal in plant–herbivore interaction? The Plant Journal 46 948–960.[CrossRef][Web of Science][Medline]
Walter A, Christ MM, Barron-Gafford GA, Grieve KA, Murthy R, Rascher U. (2005) The effect of elevated CO2 on diel leaf growth cycle, leaf carbohydrate content and canopy growth performance of Populus deltoides. Global Change Biology 11 1207–1219.
Walter A, Feil R, Schurr U. (2002a) Restriction of nyctinastic movements and application of tensile forces to leaves affects diurnal patterns of expansion growth. Functional Plant Biology 29 1247–1258.
Walter A and Schurr U. (2005) Dynamics of leaf and root growth—endogenous control versus environmental impact. Annals of Botany 95 891–900.
Walter A, Spies H, Terjung S, Küsters R, Kirchgeßner N, Schurr U. (2002b) Spatio-temporal dynamics of expansion growth in roots: automatic quantification of diurnal course and temperature response by digital image sequence processing. Journal of Experimental Botany 53 689–698.
Warneke C, Luxembourg SL, de Gouw JA, Rinne HJI, Guenther AB, Fall R. (2002) Disjunct eddy covariance measurements of oxygenated volatile organic compound fluxes from an alfalfa field before and after cutting. Journal of Geophysical Research–Atmospheres 107 doi:10.1029/2001JD000594.
Wildt J, Kobel K, Schuh-Thomas G, Heiden AC. (2003) Emissions of oxygenated volatile organic compounds from plants. Part II: emissions of saturated aldehydes. Journal of Atmospheric Chemistry 45 173–196.
Willats WGT, Orfila C, Limberg G, et al. (2001) Modulation of the degree and pattern of methyl-esterification of pectin homogalacturonan in plant cell walls. Journal of Biological Chemistry 276 19404–19413.
Willmer CM. (1983) StomataLondon Longman.
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  B. Rasulov, L. Copolovici, A. Laisk, and U. Niinemets Postillumination Isoprene Emission: In Vivo Measurements of Dimethylallyldiphosphate Pool Size and Isoprene Synthase Kinetics in Aspen Leaves Plant Physiology, March 1, 2009; 149(3): 1609 - 1618. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||