SEARCH

SEARCH BY CITATION

Keywords:

  • plant hydrocarbon emission;
  • stable carbon isotopes;
  • substrate regulation;
  • water stress;
  • temperature

ABSTRACT

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. ACKNOWLEDGMENTS
  8. REFERENCES

Isoprene is emitted from leaves of numerous plant species and has important implications for plant metabolism and atmospheric chemistry. The ability to use stored carbon (alternative carbon sources), as opposed to recently assimilated photosynthate, for isoprene production may be important as plants routinely experience photosynthetic depression in response to environmental stress. A CO2-labelling study was performed and stable isotopes of carbon were used to examine the role of alternative carbon sources in isoprene production in Populus deltoides during conditions of water stress and high leaf temperature. Isotopic fractionation during isoprene production was higher in heat- and water-stressed leaves (−8.5 and −9.3‰, respectively) than in unstressed controls (−2.5 to −3.2‰). In unstressed plants, 84–88% of the carbon in isoprene was derived from recently assimilated photosynthate. A significant shift in the isoprene carbon composition from photosynthate to alternative carbon sources was observed only under severe photosynthetic limitation (stomatal conductance < 0.05 mol m−2 s−1). The contribution of photosynthate to isoprene production decreased to 77 and 61% in heat- and water-stressed leaves, respectively. Across water- and heat-stress experiments, allocation of photosynthate was negatively correlated to the ratio of isoprene emission to photosynthesis. In water-stressed plants, the use of alternative carbon was also related to stomatal conductance. It has been proposed that isoprene emission may be regulated by substrate availability. Thus, understanding carbon partitioning to isoprene production from multiple sources is essential for building predictive models of isoprene emission.


INTRODUCTION

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. ACKNOWLEDGMENTS
  8. REFERENCES

Isoprene (2-methyl-1,3-butadiene) is emitted from leaves of numerous plant species. Understanding the physiological and environmental regulation of isoprene production has important implications for both plant metabolism and atmospheric chemistry. The function of isoprene in plants remains elusive although several hypotheses have been put forth, including thermotolerance (Singsaas et al. 1999; Sharkey, Chen & Yeh 2001), protection against oxidative stress (Loreto et al. 2001; Affek & Yakir 2002), and metabolic control (Logan, Monson & Potosnak 2000; Rosenstiel et al. 2002). Although isoprene production usually represents a loss of 1–2% of recently assimilated carbon from photosynthesis at 30 °C, carbon loss can exceed 50% under high temperature or during periods of stress when photosynthesis is depressed (Sharkey & Loreto 1993; Lerdau & Keller 1997). Isoprene also plays a major role in regulating the oxidative capacity of the troposphere and influences concentrations of important pollutants and greenhouse gases, such as ozone, carbon monoxide, and methane (reviewed in Monson & Holland 2001). Recently, it has been suggested that isoprene synthesis may be substrate limited (e.g. Rosenstiel et al. 2002) and, thus, susceptible to short- and long-term changes in chloroplast and cytosolic carbon availability. This study uses stable isotopes of carbon to examine the role of various carbon sources in isoprene production.

Isoprene is synthesized primarily in chloroplasts from dimethylallyl diphosphate (DMAPP) and isopentenyl diphosphate (IPP), which are products of the DOXP pathway (Rohmer et al. 1993; Lichtenthaler 1999). DMAPP and IPP are formed from glyceraldehyde-3-phosphate (GAP) and pyruvate, and the availability of all four compounds is influenced by processes occurring throughout the cell. Whereas GAP is initially derived in the chloroplast from the Calvin cycle, it is also imported from the cytosol, where it is an end-product of glycolysis (reviewed in Flügge 2000). Pyruvate may be synthesized in the chloroplast from glycolysis or as a by-product of ribulose bisphosphate carboxylase activity (Andrews & Kane 1991) and may also be imported from the cytosol either directly or in the form of phosphoenolpyruvate (PEP), which is then converted to pyruvate by pyruvate kinase (Fischer et al. 1997; Rosenstiel et al. 2003). Karl et al. (2002) also proposed that DMAPP precursors may be imported into the chloroplast in the form of pentose phosphates or IPP from the cytosolic mevalonic acid pathway (but see Affek & Yakir 2003). Thus, a distinction can be made between DMAPP derived from recently assimilated GAP and pyruvate in the chloroplast (photosynthetic sources) and DMAPP derived from precursors imported in various forms from processes in the cytosol (alternative sources).

There is now ample evidence that isoprene is produced from multiple carbon sources. Several studies have used isotopically labelled CO2 to examine the contribution of these various carbon pools to isoprene production. Studies using 99%13CO2 found that, at steady state, 80% of the carbon in isoprene is derived from the 13C label, indicating that roughly 20% comes from alternative sources (Sharkey et al. 1991; Delwiche & Sharkey 1993; Karl et al. 2002). Using natural abundance carbon isotope measurements, Affek & Yakir (2003) found interspecific differences in the use of alternative carbon for isoprene production, ranging between 9 and 28% incorporation of alternative carbon. These labelling studies have produced estimates of a 3–10 h turnover time of the alternative carbon source. However, additional studies have found relationships between leaf-level isoprene emission and whole-plant carbon balance, suggesting that alternative carbon sources are important on longer temporal and larger spatial scales (Funk, Jones & Lerdau 1999; Kreuzwieser et al. 2002). For example, Funk et al. (1999) found leaf emission rate to decrease by 30% following defoliation of neighbouring leaves and that leaf emission rate was positively correlated with the number of photosynthesizing leaves on the branch.

Although several studies have suggested that isoprene emission patterns are linked to isoprene synthase activity (e.g. Schnitzler et al. 1997), it has been proposed that emission can be substrate limited (via DMAPP availability) on both short- and long-time scales (Lerdau, Guenther & Monson 1997; Fall & Wildermuth 1998, Sharkey & Yeh 2001; Rosenstiel et al. 2002). Thus, understanding the partitioning of multiple carbon sources to DMAPP production may ultimately lead to the improvement of existing models of isoprene production (e.g. Guenther et al. 1993). A critical next step is to examine how dynamic these alternative carbon sources are in response to fluctuations of environmental variables that influence emission, such as light, temperature, and water availability. As a starting point, isotope labelling studies have shown that alternative carbon sources are important for maintaining isoprene production under artificial stress conditions, created by using photosynthetic inhibitors and altering CO2 concentrations to depress photosynthesis (Karl et al. 2002; Affek & Yakir 2003). Under these conditions, isoprene emission can generally continue in the absence of net CO2 uptake as long as Calvin cycle activity continues to supply substrate via photorespiration and photosynthetic electron transport continues to supply reducing power.

By varying δ13CO2, we used carbon isotope measurements to examine the role of alternative carbon in isoprene production during (1) water stress, and (2) conditions of high leaf temperature. Both prolonged water stress and elevated leaf temperature increase the ratio of isoprene emission to photosynthesis (I/P). Prolonged water stress can even result in rates of isoprene emission that exceed those of net CO2 uptake as a result of decreased photosynthesis (Tingey, Evans & Gumpertz 1981). Similarly, increasing leaf temperature will lead to a higher I/P because isoprene emission increases exponentially with temperature while photosynthesis often declines (Monson & Fall 1989; Monson et al. 1992, 1994; Sharkey & Loreto 1993). We hypothesized that an increase in I/P would lead to a change in the relative contribution of recently assimilated photosynthate and alternative carbon to isoprene production.

MATERIALS AND METHODS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. ACKNOWLEDGMENTS
  8. REFERENCES

Experimental design

Potted 2-year old Populus deltoides (clone ST109) individuals were grown in the SUNY Stony Brook, Stony Brook, NY, greenhouse with light levels ranging from 300 to 1000 µmol photon m−2 s−1 under clear sky conditions. Greenhouse temperature was regulated and ranged between 15 and 32 °C throughout the year. Plants produced new leaves throughout the year. All plants were fertilized with a slow-release fertilizer (Osmocote 19% N, 6% P, 12% K; The Scotts Company, Maryville, OH, USA) at the time of potting and were given supplemental biweekly applications of 0.3 g N (nitrate nitrogen), 0.1 g P, 0.3 g K, 0.1 g Ca, 0.04 g Mg, and micronutrients. Fully expanded leaves on the primary shoot were chosen for measurement.

The first experiment examined the effect of water stress on the partitioning of alternative carbon to isoprene production. Gas exchange measurements and isotopic analysis of isoprene and CO2 were conducted on leaves from six well-watered (group A) and seven water-stressed (group B) P. deltoides plants. Measurements were conducted before (time 1) and after (time 2) water was completely withheld from group B. Pre-stress and control plants were watered every day. The duration of water stress was variable (between 4 and 8 d) to account for differences in water loss between plants. Mid-day water potential measurements were performed on leaves from six well-watered and six water-stressed plants with a pressure chamber system (Model 610; PMS Instruments, Corvallis, OR, USA). These measurements were not performed on the plants in the experiments to avoid damage-induced effects on flux measurements, but were conducted on plants that were treated the same as those used for flux measurements. Leaf water potential ranged from −0.5 to −0.8 MPa in leaves on well-watered plants (−0.77 MPa, SE = 0.10) and −0.5 to −1.3 MPa in leaves on water-stressed plants (−0.89 MPa, SE = 0.13).

In the second experiment, gas exchange and isotopic measurements were conducted on leaves at 28 °C (low temperature, 12 leaves from 12 plants) and 38 °C (high temperature, 10 leaves from 9 plants). For these P. deltoides individuals, 24–28 °C is within the optimal temperature range for photosynthetic performance. 38 °C was chosen as the high temperature due to limitations imposed by the LI-6400 (Li-Cor Inc., Lincoln, NE, USA) thermoelectric devices and infra-red gas analyser (IRGA) reliability. All plants used in the heat-stress experiment were well-watered.

Gas exchange

Photosynthetic and isoprene emission rates were measured using two open path LI-6400 portable photosynthesis systems and a Photovac Voyager gas chromatograph (GC) with a photo-ionization detector (Perkin-Elmer, Norwalk, CT, USA) as described in Funk et al. (2003). The LI-6400 has a temperature- and light-controlled leaf chamber, which houses IRGAs. Flow rates through the cuvette and GC ballast line varied between 75 and 350 µmol s−1. All measurements were taken with a variable intensity red/blue light emitting diode, with cuvette light levels at 1500 µmol photon m−2 s−1 and peak irradiance at 665 and 470 nm. During the water-stress experiment, dual thermoelectric devices maintained leaf temperature at 28 °C. A desiccant trap located prior to the leaf chamber held humidity inside the chamber at 30 ± 10%.

The CO2 concentration of air entering the leaf chamber was maintained at 400 p.p.m.v. by diluting pure CO2 through a mass flow control system inside the LI-6400. Leaves were fed CO2 with three distinct isotopic signatures (−6.1, −43.0, and −105.8‰; V-PDB-CO2). Due to fractionation associated with leaks and incomplete air flushing in the LI-6400, multiple chamber blank measurements were conducted to accurately assess δ13CO2 entering the leaf chamber, and CO2 sources were adjusted (−6.8, −43.5, −104.0‰). To minimize potential variability associated with different leaves, measurements with the three CO2 sources were taken on different 6 cm2 sections of the same leaf. Leaf sections sampled at a common δ13CO2 showed no significant differences in δ13C of isoprene (J. Funk, unpublished results). Leaf sections were allowed to equilibrate inside the leaf chamber for 20–30 min prior to measurement.

Sample air was routed from the cuvette into a 1.75-L Teflon ballast line, which was flushed twice with sample air before measurement. Air was drawn from this line into the 1 mL sample loop within the GC at 100 mL min−1 for 10 s. We used a 15-m Quadrex 007–1 methyl silicone phase capillary column (12 µm interior diameter, 0.32 mm coating thickness), maintained at 50 °C, with UHP nitrogen as a carrier gas. The detector output was linear across the range of isoprene concentrations used, with an intercept through zero. Minimum detection was 3 p.p.b.v. and detector reproducibility was within 10%. Two-point calibrations were made daily with dilutions from a 97.9 p.p.m.v. mix of isoprene in air (Scott-Marrin, Riverside, CA, USA).

Isotopic analysis of isoprene and CO2

Air was diverted out of the leaf chamber to two 150 mL sample bottles in series via a dry ice trap to remove water. Flow through the collection bottles was maintained above 70 mL min−1 for 20 min. δ13C of isoprene was measured using a continuous flow isotope ratio mass spectrometer (IRMS) (Finnigan Delta Plus; Finnigan MAT, Bremen, Germany) with a modified pre-concentration GC/combustion interface similar to that described in Mak & Yang (1998). Sample air was cryogenically trapped on the pre-concentration unit and the concentrated sample was loaded onto the GC column (0.32 mm × 30 m; HP-5) at a flow rate of 3.7 mL min−1 at −35 °C. Isoprene was then combusted to CO2 at 900 °C. CO2 and isoprene-derived CO2 were then sent to the IRMS separated by 250 s. A 200 p.p.b.v. isoprene standard diluted from a 97.9 p.p.m.v. mix of isoprene in air (Scott-Marrin) was run daily. Repeated sampling of this standard (−19.94‰; V-PDB) generated a 95% confidence limit of 0.90‰. Isotope data are expressed with conventional δ notation:

  • image

where R is the ratio of mass 45 (13CO2) to mass 44 for sample and standard (V-PDB-CO2).

The CO2 was isolated from sample air on a vacuum extraction line with dry ice and liquid nitrogen traps. Once purified, CO2 samples were run following a conventional dual-inlet approach on the IRMS described above. Photosynthetic discrimination (Δps) was calculated by comparing δ13CO2 of air entering and exiting the leaf chamber as in Evans et al. (1986):

  • image

where δo and δe represent the isotopic composition of CO2 exiting and entering the leaf chamber, respectively, and

  • image

where ce and co are the CO2 concentrations in air entering and exiting the leaf chamber.

When feeding leaves isotopically depleted CO2, discrimination calculations using the above equations can overestimate Δps (Gillon et al. 1998). This occurs because δ13C of CO2 from respiration and photorespiration diffusing out of the leaf can be isotopically enriched relative to that of source CO2. This phenomenon leads to larger changes in δ13C between CO2 entering and exiting the leaf chamber and artificially high Δps values. We addressed this issue by only calculating Δps when −6.12‰ CO2 was used and we assumed that Δps was constant across all leaf sections.

Leaf tissue analyses

Leaves used for leaf water potential measurements from six well-watered and six water-stressed plants were frozen in liquid nitrogen, dried at 70 °C for 48 h, and ground. The isotopic composition of leaves was determined by continuous-flow IRMS, consisting of an elemental analyser (Model NC2500; Carlo Erba, Milan, Italy) interfaced to an IRMS system (Finnigan Delta Plus). Cabbage (−27.15‰, SD = 0.08) and methionine (−25.38‰, SD = 0.11) were used as standards for precision and linearity, respectively. Samples were measured against CO2 reference gas (−42.39‰; PDB-CO2). Leaf δ13C averaged −30.5‰ (SE = 0.23) and −25.5‰ (SE = 0.36) in well-watered and water-stressed plants, respectively.

Statistical analysis

Multivariate repeated measures analysis of variance was used to assess the effect of water stress on δ13C and gas exchange variables (proc glm, SAS 6.12; SAS Institute Inc., Cary, NC, USA). P-values reported are those from the group (A,B) by time interaction term. Differences in δ13C and gas exchange variables due to leaf temperature were examined with t-tests (Statistica 5.1, Statsoft, Tulsa, OK, USA). Data that violated the anova assumptions of normality and homogeneity of variance were rank transformed. Corrections for multiple comparisons did not alter the significance of any analysis.

Pearson product–moment correlation coefficients were generated to evaluate the linear association between δ13C of photosynthate and δ13C of isoprene (Biomstat 3.30j; Applied Biostatistics, Port Jefferson, NY, USA). Slopes and associated confidence intervals were derived from the principal axis, which corresponds to the major axis regression line found in model I regression. Differences between slopes were evaluated with tests of parallelism (Statistica 5.1). Measurement error for all observations were excluded because of the large range over which variables were examined.

RESULTS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. ACKNOWLEDGMENTS
  8. REFERENCES

Leaf gas exchange

Isoprene emission rates in water-stressed plants (group B) increased slightly relative to well-watered, control plants (group A) (Table 1). In contrast, photosynthetic rate, stomatal conductance, and intercellular CO2 concentration (ci) declined in water-stressed plants relative to controls. In stressed plants, photosynthesis and conductance fell to less than half of pre-stress values, leading to a four-fold increase in I/P (Table 1).

Table 1.  Gas exchange results from the water-stress experiment
 Time 1Time 2P-value (F-value)
ABAB
  1. Individuals in group A (n = 6) were watered daily while water was completely withheld from individuals in group B (n = 7) after measurement in Time 1 until re-measurement in Time 2. See ‘Methods: Experimental design’ for description of water-stress treatment. Numbers presented are means ± standard errors. The statistical significance of water stress on variables is represented by the time by group interaction term from multivariate repeated measures anova. We report P-values followed by F-values in parentheses.

Isoprene emission (nmol m−2 s−1)40.4 ± 3.437.6 ± 2.033.6 ± 2.548.8 ± 7.0  0.002 (15.71)
Photosynthetic rate (µmol m−2 s−1)18.6 ± 0.818.2 ± 0.318.8 ± 0.9 6.8 ± 1.2< 0.0001 (68.43)
Stomatal conductance (mol m−2 s−1)0.41 ± 0.040.54 ± 0.070.40 ± 0.040.06 ± 0.01< 0.0001 (180.99)
Intercellular CO2 (µL L−1) 201 ± 6 215 ± 4 192 ± 7 156 ± 7< 0.0001 (185.44)
% of photosynthesis as isoprene emission0.22 ± 0.020.19 ± 0.020.18 ± 0.020.79 ± 0.13< 0.001 (20.68)

As expected, isoprene emission increased with leaf temperature (Table 2). Leaves measured at 38 °C displayed significantly lower assimilation rates than leaves measured at 28 °C, despite similar stomatal conductance and higher ci. As in the water-stress experiment, the decoupling of isoprene emission and photosynthesis led to a four-fold increase in I/P.

Table 2.  Gas exchange results from the heat-stress experiment (n = 12 leaves at 28 °C, n= 10 leaves at 38 °C)
 Temperature (°C) P-value (F-value)
Low (28)High (38)
  1. Numbers presented are means ± standard errors. P-values (with F-values) were generated with t-tests.

Isoprene emission (nmol m−2 s−1)30.2 ± 1.581.6 ± 5.8< 0.0001 (86.02)
Photosynthetic rate (µmol m−2 s−1)18.4 ± 0.414.4 ± 1.1< 0.01 (13.18)
Stomatal conductance (mol m−2 s−1)0.44 ± 0.030.38 ± 0.06  0.34 (0.97)
Intercellular CO2 (µL L−1) 207 ± 4 247 ± 6< 0.0001 (32.69)
% of photosynthesis as isoprene emission0.22 ± 0.010.60 ± 0.07< 0.0001 (58.54)

Carbon sources for isoprene production

To examine the relative contribution of recently assimilated photosynthate to isoprene production, we plotted measured δ13C of isoprene against calculated δ13C of photosynthate as in Affek & Yakir (2003) (Fig. 1, Table 3). A slope of 1.0 represents a perfect correlation between the variables and, thus, 100% incorporation of photosynthate into isoprene. This approach assumes that partitioning of multiple carbon sources and isotopic discrimination during photosynthesis and isoprene production was similar across the δ13C range of CO2 sources used. Fractionation during isoprene production was calculated from the offset between δ13C of isoprene and photosynthate (Affek & Yakir 2003) at the value of bulk leaf carbon (−30.5‰ in well-watered leaves, −25.5‰ in water-stressed leaves). When photosynthate and bulk leaf carbon are at the same value, the difference in δ13C between isoprene and photosynthate reflects fractionation during isoprene production. This method assumes that the δ13C of alternative carbon is equivalent to bulk leaf carbon.

image

Figure 1. The relationship between the isotopic signature of carbon in isoprene and photosynthate in (a) leaves from well-watered (n = 6; r = 1.0; •) and water-stressed (n = 7; r = 0.97; ○) plants and (b) leaves at 28 °C (n = 12; r = 1.0; ▪) and 38 °C (n = 10; r = 0.98; □) using three isotopically distinct CO2 sources (−6.1, −43.0, −105.8‰). δ13C of isoprene was measured directly and δ13C of photosyntate was calculated using the method of Evans et al. (1986).

Download figure to PowerPoint

Table 3.  The percentage of photosynthate-derived carbon in isoprene and isotopic discrimination during isoprene production in water-stress and heat-stress experiments
Water stressABP-value (F-value)
Time 1
 % C from photosynthate84.4 ± 3.383.7 ± 2.8 0.73 (0.12)
 Fractionation (‰)−2.5−3.2 
Time 2
 % C from photosynthate88.1 ± 2.862.0 ± 7.2<0.0001 (45.34)
 Fractionation (‰)−3.2−9.3 
Heat stress28 °C38 °CP-value (F-value)
  1. Water-stress treatments are described in Table 1. The percentage of photosynthate-derived carbon in isoprene was calculated from correlation analysis of δ13C of isoprene and δ13C of photosynthate. Numbers presented are means ± 95% confidence limits. P-values (with F-values) are from tests of parallelism between slopes. Fractionation values were calculated from the offset between δ13C of isoprene and bulk leaf carbon.

% C from photosynthate86.6 ± 2.377.1 ± 5.4<0.001 (14.41)
Fractionation (‰)−2.9−8.5 

In well-watered plants, 84–88% of the carbon in isoprene was derived from photosynthate (Table 3, Fig. 1). This percentage dropped significantly in water-stressed plants to 61%. The substantial variation in the relative contribution of photosynthate to isoprene production in water-stressed plants (Fig. 1a) was partly explained by the degree of stress (Fig. 2). Two moderately stressed plants (stomatal conductance 0.10–0.12 mol m−2 s−1) continued to incorporate a relatively high percentage of photosynthate-derived carbon into isoprene (71–81%). Only in severely water-stressed plants (stomatal conductance < 0.05 mol m−2 s−1) did the percentage of photosynthate-derived carbon in isoprene fall below 70%. However, despite large reductions in carbon assimilation, the most stressed individuals continued to acquire roughly half of the carbon needed for isoprene production from photosynthate (Fig. 2). Fractionation during isoprene production was three times higher in leaves of water-stressed plants relative to well-watered plants (Table 3).

image

Figure 2. The relationship between stomatal conductance and the percentage of photosynthate-derived carbon in isoprene in well-watered (n = 6; •) and water-stressed (n = 7; ○) plants at Time 2 (after stress).

Download figure to PowerPoint

The percentage of photosynthate-derived carbon in isoprene was significantly smaller in leaves at high temperature (77%) relative to those measured at 28 °C (87%) (Table 3, Fig. 1b). However, the percentage was larger than that in water-stressed plants. As in the water-stress experiment, fractionation during isoprene production was roughly three times higher in heat-stressed leaves relative to leaves measured at lower temperature (Table 3). Across water- and heat-stressed leaves, the percentage of photosynthate-derived carbon in isoprene was negatively correlated with I/P (r = −0.84, P < 0.0001; Fig. 3).

image

Figure 3. The relationship between the ratio of isoprene emission to photosynthesis (I/P) and the percentage of photosynthate-derived carbon in isoprene in unstressed (n = 18, •), heat-stressed (n = 10, ○), and water-stressed (n = 7, ▴) leaves.

Download figure to PowerPoint

DISCUSSION

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. ACKNOWLEDGMENTS
  8. REFERENCES

Fractionation during isoprene production

The carbon isotope signature of isoprene is determined by: (1) discrimination during photosynthesis (Δps) and isoprene synthesis (Δiso); (2) δ13CO2; and (3) δ13C of alternative carbon sources feeding DMAPP production. In this study, Δiso was small in unstressed plants (−2.5 to −3.2‰) and larger in heat- (−8.5‰) and water-stressed (−9.3‰) leaves, relative to bulk leaf carbon. Δiso has only been examined in a few studies. Isoprene emitted from red oak and velvet bean was found to be isotopically depleted relative to recently assimilated carbon (photosynthate) by 2.8 and 2.6‰, respectively (Sharkey et al. 1991; Rudolph et al. 2003). However, Affek & Yakir (2003) found a large range in Δiso, from −4.0 to −10.6‰ across different species of terrestrial plants. Schouten et al. (1998) also found substantial variation in isotopic discrimination during phytol production in different species of marine algae (+0.9 to −5.3‰; relative to bulk cell material). A potential error in Δiso in this study and in Affek & Yakir (2003) arises from the assumption that δ13C of alternative carbon is equivalent to bulk leaf carbon.

Affek & Yakir (2003) propose that the main point of isotopic discrimination during isoprene production is associated with the decarboxylation of pyruvate. In this experiment, heat stress may have affected this process, leading to an increase in Δiso. DeNiro & Epstein (1977) found temperature-dependent kinetic isotope effects of pyruvate decarboxylase between 15 and 35 °C when the decarboxylation reaction was not run to completion. The degree to which this reaction is carried to completion, which decreases the potential for fractionation, depends on pyruvate availability, mediated by demand from other reactions. However, Rudolph et al. (2003) found little temperature dependence in δ13C of isoprene over a 15 °C temperature range. Alternatively, stress could alter the relative contribution of alternative carbon sources with differing δ13C. For example, Kreuzwieser et al. (2002) found increased incorporation of glucose as an alternative carbon source for isoprene production under high temperature. Furthermore, it has been suggested that fractionation may occur during metabolite transport from the cytosol to the chloroplast (Brugnoli & Farquhar 2000). Fractionation during carbon metabolism, including the synthesis of secondary metabolites, requires further research.

Carbon sources for isoprene production

Our data support previous findings that photosynthate is the primary source for DMAPP (e.g. Sharkey et al. 1991). In unstressed plants, 84–88% of the carbon in isoprene was derived from photosynthate. These results are comparable with the 72–91% range found in another study using natural isotope abundance measurements (Affek & Yakir 2003). Studies using 99%13CO2 (Delwiche & Sharkey 1993; Karl et al. 2002) observe only 80% labelling of isoprene by photosynthate carbon, which parallels labelling patterns of phosphoglyceric acid (PGA) (Canvin 1979), an indicator of carbon turnover in the Calvin cycle. Currently, it is unclear why natural abundance studies find > 80% incorporation of photosynthate into isoprene. The incorporation of CO2 into PGA, isoprene, and other photosynthetic products may vary between species, with plant age, and across environmental conditions (e.g. Canvin 1979).

Across water- and heat-stress experiments, a significant contribution of alternative carbon to isoprene production (>30%) was observed only under severe photosynthetic limitation (stomatal conductance < 0.05 mol m−2 s−1; Fig. 2). Variation in the use of alternative carbon was a function of the ratio of isoprene emission to photosynthesis (I/P; Fig. 3). In both stress experiments, increased I/P was attributable to elevated rates of isoprene emission and reduced rates of photosynthesis. It must be noted that, although I/P increased by a factor of 4 in stressed leaves relative to controls, the amount of carbon assimilated during photosynthesis was roughly 150 times greater than that allocated to isoprene production. Thus, the correlation may not reflect a limitation of photosynthate in stressed leaves.

In water-stressed plants, carbon source partitioning was also related to stomatal conductance (Fig. 2). Although heat-stressed plants displayed decreased photosynthetic rates, stomatal closure did not occur, which corresponds with recent documentation that photosynthesis can decrease independently of stomatal activity through changes in ATP synthesis (Tezara et al. 1999). Thus stomatal conductance does not appear to be a reliable parameter for assessing carbon source partitioning to isoprene production (or production rates) on short time scales.

Whereas the heat-stress experiment exposed individual leaves to high temperature for 20- to 30-min intervals, whole plants were subjected to water stress for 4–8 d. Thus, both recently fixed and stored carbon pools were affected by stress in the water-stress experiment whereas only recently fixed carbon pools were affected in the short heat-stress experiment. It would seem that less alternative carbon would be available for isoprene production in the water-stressed plants as stored carbon pools are depleted over several days. Alternatively, plants undergoing stress for multiple days may increase carbon import from root and stem storage into leaves, which could account for the greater incorporation of alternative carbon into isoprene in water-stressed plants. Recently assimilated photosynthate accounted for 62% of the carbon in isoprene in water-stressed plants and 77% of the carbon in isoprene in heat-stressed plants. At this point, it is unclear how the type and duration of stress influence isoprene emission rate and the allocation of various carbon sources to isoprene production.

Implications for predictive models of isoprene emission

Understanding the physiological regulation of isoprene emission from plants has challenged scientists for decades (e.g. Rasmussen & Went 1965, Sanadze & Kalandadze 1966, Tingey et al. 1979). Regulation on very short-time scales is dominated by changes in light and temperature on the order of minutes (Guenther et al. 1993). However, our knowledge of the mechanisms by which physiological and environmental factors regulate the standardized emission rate of isoprene (SER, emission at a standard light and temperature) over larger temporal (e.g. light and temperature growth environment) and spatial (e.g. resource availability) scales is incomplete and often species or system specific. As isoprene SER is a fundamental parameter in emission models (e.g. Guenther et al. 1993), understanding the response of isoprene SER to environmental and physiological parameters will improve regional and global predictions of isoprene emission.

The gas exchange results for our water-stress experiment contrast with those from previous studies that have found that isoprene SER is not affected by water stress (Tingey et al. 1981; Guenther et al. 1999) or is mildly depressed (Sharkey & Loreto 1993; Fang, Monson & Cowling 1996; Lerdau et al. 1997; Brüggemann & Schnitzler 2002). The lack of a general emission pattern in response to water stress across many species and study protocols (e.g. plant age, growth temperature, duration of stress) suggests that many variables may influence isoprene production, possibly through changes in substrate availability and enzymatic activity.

The dependence on alternative carbon sources to sustain isoprene production under conditions of low assimilation suggests that some measure of leaf or whole-plant carbon status could be used when modelling emission during drought or prolonged temperature stress. Alternative sources may play a major role in response to water stress on both long (e.g. low soil water potential during periods of drought) and short (e.g. mid-day stomatal closure associated with increased vapor pressure deficit) time scales. Furthermore, an increased contribution of alternative carbon to isoprene production may explain the increased isoprene SER during afternoon measurements, which coincides with decreases in photosynthesis (Steinbrecher et al. 1997; Sharkey et al. 1999; Funk et al. 2003). Understanding the contribution of alternative carbon sources to isoprene production is paramount if emission is subject to substrate regulation (e.g. Rosenstiel et al. 2002). Results from this and other studies (Funk et al. 1999; Kreuzwieser et al. 2002) suggest that future research should address substrate regulation on cellular to whole-plant levels.

ACKNOWLEDGMENTS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. ACKNOWLEDGMENTS
  8. REFERENCES

We thank Laura Cottrell, Art Kasson, Ted Sandomenico, Wenbo Yang, and Lee Ferguson for laboratory and technical assistance, Mike Axelrod and John Klumpp for greenhouse assistance, Jessica Gurevitch, Peter Harley, and Dianna Padilla for comments on the manuscript, and Darren Sandquist, Laura Hyatt, and Tom Sharkey for comments on an earlier version of the manuscript. This work was funded by an NSF dissertation improvement grant (DEB-0206071) and EPA and NSF predoctoral fellowships to J.L.F., NSF (DEB-9552658) and NASA (NAGW-5249) research grants to M.T.L.

REFERENCES

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. ACKNOWLEDGMENTS
  8. REFERENCES
  • Affek H.P. & Yakir D. (2002) Protection by isoprene against singlet oxygen in leaves. Plant Physiology 0, 269277.
  • Affek H.P. & Yakir D. (2003) Natural abundance carbon isotope composition of isoprene reflects incomplete coupling between isoprene synthesis and photosynthetic carbon flow. Plant Physiology 131, 17271736.
  • Andrews T.J. & Kane H.J. (1991) Pyruvate is a by-product of catalysis by ribulosebisphosphate carboxylase/oxygenase. Journal of Biological Chemistry 266, 94479452.
  • Brüggemann N. & Schnitzler J.-P. (2002) Comparison of isoprene emission, intercellular isoprene concentration and photosynthetic performance in water-limited oak (Quercus pubescens Willd. & Quercus robur L.) saplings. Plant Biology 4, 456463.
  • Brugnoli E. & Farquhar G.D. (2000) Photosynthetic fractionation of carbon isotopes. In: Photosynthesis: Physiology and Metabolism (eds R.C.Leegood, T.D.Sharkey & S.Von Caemmerer), pp. 399434. Kluwer Academic Publishers, Dordrecht, The Netherlands.
  • Canvin D.T. (1979) Photorespiration: comparison between C3 and C4 plants. In: Encyclopedia of Plant Physiology, Vol. 6: Photosynthesis II (eds M.Gibbs & E.Latzko), pp. 368396. Springer-Verlag, Berlin, Germany.
  • Delwiche C. & Sharkey T. (1993) Rapid appearance of 13C in biogenic isoprene when 13CO2 is fed to intact leaves. Plant, Cell and Environment 16, 587591.
  • DeNiro M.J. & Epstein S. (1977) Mechanism of carbon isotope fractionation associated with lipid synthesis. Science 197, 261263.
  • Evans J.R., Sharkey T.D., Berry J.A. & Farquhar G.D. (1986) Carbon isotope discrimination measured concurrently with gas exchange to investigate CO2 diffusion in leaves of higher plants. Australian Journal of Plant Physiology 13, 281292.
  • Fall R. & Wildermuth M. (1998) Isoprene synthase: from biochemical mechanism to emission algorithm. Journal of Geophysical Research 103, 2559925609.
  • Fang C., Monson R. & Cowling E. (1996) Isoprene emission, photosynthesis, and growth in sweetgum (Liquidambar styraciflua) seedlings exposed to short- and long-term drying cycles. Tree Physiology 16, 441446.
  • Fischer K., Kammerer B., Gutensohn M., Arbinger B., Weber A., Hausler R.E. & Flügge U.-I. (1997) A new class of plastidic phosphate translocators: a putative link between primary and secondary metabolism by the phosphoenolpyruvate/phosphate antiporter. Plant Cell 9, 453462.
  • Flügge U.-I. (2000) Metabolite transport across the chloroplast envelope of C3-plants. In: Photosynthesis: Physiology and Metabolism (eds R.C.Leegood, T.D.Sharkey & S.Von Caemmerer), pp. 137152. Kluwer Academic Publishers, Dordrecht, The Netherlands.
  • Funk J.L., Jones C.G., Baker C.J., Fuller H.M., Giardina C.P. & Lerdau M.T. (2003) Diurnal variation in the basal emission rate of isoprene. Ecological Applications 13, 169178.
  • Funk J.L., Jones C.G. & Lerdau M.T. (1999) Defoliation effects on isoprene emission from Populus deltoides. Oecologia 118, 333339.
  • Gillon J.S., Borland A.M., Harwood K.G., Roberts A., Broadmeadow M.S.J. & Griffiths H. (1998) Carbon isotope discrimination in terrestrial plants: carboxylations and decarboxylations. In: Stable Isotopes: Integration of Biological, Ecological and Geochemical Processes (ed. H.Griffiths), pp. 111131. BIOS Scientific Publishers Ltd, Oxford, UK.
  • Guenther A., Archer S., Greenberg J., Harley P., Helmig D., Klinger L., Vierling L., Wildermuth M., Zimmerman P. & Zitzer S. (1999) Biogenic hydrocarbon emissions and landcover/climate change in a subtropical savanna. Physics and Chemistry of the Earth (B) 24, 659667.
  • Guenther A.B., Zimmerman P.R., Harley P.C., Monson R.K. & Fall R. (1993) Isoprene and monoterpene emission rate variability: Model evaluations and sensitivity analyses. Journal of Geophysical Research 98, 1260912617.
  • Karl T., Fall R., Rosenstiel T.N., Prazeller P., Larsen B., Seufert G. & Lindinger W. (2002) On-line analysis of the 13CO2 labeling of leaf isoprene suggests multiple subcellular origins of isoprene precursors. Planta 215, 894905.
  • Kreuzwieser J., Graus M., Wisthaler A., Hansel A., Rennenberg H. & Schnitzler J.-P. (2002) Xylem-transported glucose as an additional carbon source for leaf isoprene formation in Quercus robur. New Phytologist 156, 171178.
  • Lerdau M. & Keller M. (1997) Controls on isoprene emission from trees in a subtropical dry forest. Plant, Cell and Environment 20, 569578.
  • Lerdau M., Guenther A. & Monson R. (1997) Plant production and emission of volatile organic compounds. Bioscience 47, 373383.
  • Lichtenthaler H.K. (1999) The 1-deoxy-d-xylulose-5-phosphate pathway of isoprenoid biosynthesis in plants. Annual Review of Plant Physiology and Molecular Biology 50, 4765.
  • Logan B.A., Monson R.K. & Potosnak M.J. (2000) Biochemistry and physiology of foliar isoprene production. Trends in Plant Science 5, 477481.
  • Loreto F., Mannozzi M., Maris C., Nascetti P., Ferranti F. & Pasqualini S. (2001) Ozone quenching properties of isoprene and its antioxidant role in leaves. Plant Physiology 126, 9931000.
  • Mak J.E. & Yang W. (1998) Technique for analysis of air samples for 13C and 18O in carbon monoxide via continuous-flow isotope ratio mass spectrometry. Analytical Chemistry 70, 51595161.
  • Monson R. & Fall R. (1989) Isoprene emission from aspen leaves: influence of environment and relation to photosynthesis and photorespiration. Plant Physiology 90, 267274.
  • Monson R.K. & Holland E.A. (2001) Biospheric trace gas fluxes and their control over tropospheric chemistry. Annual Review of Ecology and Systematics 32, 547576.
  • Monson R., Harley P., Litvak M., Wildermuth M., Guenther A., Zimmerman P. & Fall R. (1994) Environmental and developmental controls over the seasonal pattern of isoprene emission from aspen leaves. Oecologia 99, 260270.
  • Monson R., Jaeger C., Adams W.I., Drigger E., Silver G. & Fall R. (1992) Relationships among isoprene emission rate, photosynthesis, and isoprene synthase activity as influenced by temperature. Plant Physiology 98, 11751180.
  • Rasmussen R.A. & Went F.W. (1965) Volatile organic material of plant origin in the atmosphere. Proceedings of the National Academy of Science USA 53, 215220.
  • Rohmer M., Knani M., Simonin P., Sutter B. & Sahm H. (1993) Isoprenoid biosynthesis in bacteria – a novel pathway for the early steps leading to isopentenyl diphosphate. Biochemical Journal 295, 517524.
  • Rosenstiel T.N., Fisher A.J., Fall R. & Monson R.K. (2002) Differential accumulation of dimethylallyl diphosphate in leaves and needles of isoprene- and methylbutenol-emitting and nonemitting species. Plant Physiology 129, 12761284.
  • Rosenstiel T.N., Potosnak M.J., Griffin K.L., Fall R. & Monson R.K. (2003) Increased CO2 uncouples growth from isoprene emission in an agriforest ecosystem. Nature 421, 256259.
  • Rudolph J., Anderson R.S., Czapiewski K.V., Czuba E., Ernst D., Gillespie T., Huang L., Rigby C. & Thompson A.E. (2003) The stable carbon isotope ratio of biogenic emissions of isoprene and the potential use of stable isotope ratio measurements to study photochemical processing of isoprene in the atmosphere. Journal of Atmospheric Chemistry 44, 3955.
  • Sanadze G.A. & Kalandadze A.N. (1966) Light and temperature curves of the evolution of C5H8. Soviet Plant Physiology 13, 411413.
  • Schnitzler J.-P., Lehning A. & Steinbrecher R. (1997) Seasonal pattern of isoprene synthase activity in Quercus robor leaves and its significance for modeling isoprene emission rates. Botanica Acta 110, 240243.
  • Schouten S., Breteler W.C.M.K., Blokker P., Schogt N., Rupstra W.I.C., Grice K., Baas M. & Damste J.S.S. (1998) Biosynthetic effects on the stable carbon isotopic compositions of algal lipids: implications for deciphering the carbon isotopic biomarker record. Geochimica et Cosmochimica Acta 62, 13971406.
  • Sharkey T.D. & Loreto F. (1993) Water stress, temperature, and light effects on the capacity for isoprene emission and photosynthesis of kudzu leaves. Oecologia 95, 328333.
  • Sharkey T.D. & Yeh S. (2001) Isoprene emission from plants. Annual Review of Plant Physiology and Plant Molecular Biology 52, 407436.
  • Sharkey T.D., Chen X. & Yeh S. (2001) Isoprene increases thermotolerance of fosmidomycin-fed leaves. Plant Physiology 125, 20012006.
  • Sharkey T.D., Loreto F., Delwiche C.F. & Treichel I.W. (1991) Fractionation of carbon isotopes during biogenesis of atmospheric isoprene. Plant Physiology 97, 463466.
  • Sharkey T.D., Singsaas E., Lerdau M. & Geron C. (1999) Weather effects on isoprene emission capacity and applications in emissions algorithms. Ecological Applications 9, 11321137.
  • Singsaas E.L., Laporte M.M., Shi J.-Z., Monson R.K., Bowling D.R., Johnson K., Lerdau M., Jasentuliytana A. & Sharkey T.D. (1999) Kinetics of leaf temperature fluctuation affect isoprene emission from red oak (Quercus rubra) leaves. Tree Physiology 19, 917924.
  • Steinbrecher R., Hauff K., Rabong R. & Steinbrecher J. (1997) Isoprenoid emission of oak species typical for the Mediterranean area: Source strength and controlling variables. Atmospheric Environment 31, 7988.
  • Tezara W., Mitchell V.J., Driscoll S.D. & Lawlor D.W. (1999) Water stress inhibits plant photosynthesis by decreasing coupling factor and ATP. Nature 401, 914917.
  • Tingey D.T., Evans R. & Gumpertz M. (1981) Effects of environmental conditions on isoprene emission from live oak. Planta 152, 656570.
  • Tingey D.T., Manning M., Grothaus L.C. & Burns W.F. (1979) The influence of light and temperature on isoprene emission rates from live oak. Physiologia Plantarum 47, 112118.