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Acetone emission from conifer buds
Phytochemistry, Vol. 34, No. 4, pp. 99-994, Printedin Great Britain
1993
ACETONE
0031~9422/93 s6.00+0.00 0 1993 PerganmnPress Ltd
EMISSION FROM CONIFER ROBERT
Cooperative
Institute
C.
MACDONALD
BUDS
and RAY FALL
for Research in Environmental Sciences and Department of Chemistry and Biochemistry, University of Colorado, Campus Box 216, Boulder, CO 80309-0216, U.S.A. (Received
Key Word Index-Abies tolerance; dormancy.
in revised form 5 April 1993)
concolor; Pinaceae; gymnosperm;
conifer; emission; ketone; acetone; bud; cold
Abstract-Acetone was detected in the headspace of excised vegetative buds of all seven conifers examined. Acetone emission rates of 7.4 + 1.5 pg hr- ’ g- 1fr. wt were measured from intact buds of Abies concolor during a growth chamber experiment. Little or no acetone emission was observed from the leaves of conifers or from any angiosperm vegetative tissue tested. Acetone headspace concentrations from buds of three conifer species were measured over two growing seasons at a forest site. Acetone concentrations were highest in winter and dropped precipitously with the onset of budbreak, suggesting that acetone may play a role in cold tolerance in gymnosperms.
INTRODUCTION
Plants are known to emit a large number of volatile compounds, some of which, such as ethylene, ethanol and monoterpenes, are known to play important physiological roles [I -31. Recently, relatively high concentrations of acetone have been measured in the atmosphere of rural pine forests [4-61. The presence of substantial amounts of acetone in rural air raised the question of whether plants could be a significant source of acetone. Isidorov et al. [7] list acetone as being produced by 22 species of plants, as determined by GC-mass spectrometry. Unfortunately, no detailed data were presented; thus, the quantities of acetone emitted and the source of acetone production within the plants are unclear. We could find no reports on the biology of acetone emission from vegetative tissues. There have been several reports in the literature of acetone production by germinating seeds [S-11], possibly in association with lipid metabolism [l 11. Much of the work with acetone production by seeds involved elevated incubation temperatures [9-l 11, leading to the possibility of artifacts such as the thermal decomposition of acetoacetate to acetone. The purpose of this research was to determine whether plants are capable of emitting sufficient amounts of acetone to account for that detected in rural air. We are also interested in determining the role of acetone emissions in plants, and in determining how acetone is produced.
UESULTS
In order to determine whether vegetative tissues were capable of emitting significant quantities of acetone, leaves and buds from trees on the University of Colorado
campus were sampled in a species survey. Little or no acetone was detected in the leaf emissions of any species sampled (Table 1). However, acetone was emitted by the excised vegetative buds of every gymnosperm species studied in headspace concentrations of up to 25.5 ngml-i (Table l), and as high as 200 ngml-’ in later experiments (Fig. 1). Little or no acetone was emitted from the buds of angiosperms (Table 1). Acetone production was not significantly altered by surface sterilization of buds of Abies concolor (P=O.2578, Wilcoxon signed rank test). Acetone production in excised candles of Pinus Jexilis exhibited a Q10 of 2.17kO.54 (Q,e is the factor by which a reaction rate increases with a 10” increase in temperature). To examine whether intact tissues emit acetone, and to determine the effect of budbreak under isothermal conditions, emission rates from intact buds and needles of A. concolor were measured in a growth chamber experiment. At 25”, acetone emission rates were 7.4 f 1.5 pg hr- ’ g- i fr. wt before budbreak, declining to 1.0 +0.2 pghr- 1g - 1fr. wt by the needles produced from these buds (n = 4). Similarly, the concentration of a putative precursor of acetone, acetoacetate, also declined from 46.4k9.2 pgg-l fr. wt in buds to 0.0k4.3 pgg-i fr. wt in the newly-produced needles. To verify the uniqueness of acetone emission to gymnosperm buds in the field, and to more precisely determine the pattern of acetone emission during budbreak, acetone emission from the buds and leaves of Pinus contorta, Abies lasiocarpa, Picea engelmannii and Populus tremuloides was measured over the course of budbreak at a pristine alpine site at the University of Colorado Mountain Research Station in 1991. Substantial production of acetone by the buds of each gymnosperm species was observed, with production declining precipitously 991
R. C. MACDONALDand R. FALL
992
Table 1. Headspace acetone concentrations from leaves and buds of trees Headspace acetone Leaves
Species
Buds (ngml-‘)
Gymnosperms Abies concolor (Gord. & Glend.) Lindl. ex Hildbr. A. lusiocurpa (Hook) Nutt. Picea glauca (Moench) Voss P. pungens Engelm. P. engelmannii Engelm. Pseudotsuga menziesii (Mirb.) Franc0 Pinus contorta Loud.
0.8&0.4 0.2 f 0.2 nd nd _
22.6 + 6.6 25.5 5 2.4 3.8 +0.2 6.8 +0.7 8.2k2.7 2.0 + 0.9 19.3* 7.5
Angiosperms Acer saccharinum L. A. rubrum L. Quercus rubra L. Tilia americana L. Populus tremuloides Michx.
*Data are meanfs.e.
1.1+0.2 nd nd tr nd
tr tr nd
of headspace acetone concentrations produced by three samples
for each species. tr, Trace amounts present; nd, not detectable.
buds increased during the winter, reaching a midwinter peak before declining to values similar to those obtained during the spring of 1991. DISCUSSION
JJASONDJFMAMJJA 1991
I
1992
Fig. 1. Variation in acetone emission from conifer buds with time. Data are mean and s.e. of three samples per species. Arrows indicate where budbreak occurred for all three species.
upon leafout (Fig. 1, inset). An exception was P.contorta, in which some residual acetone emission occurred during the candle stage of development. Acetone production was not detected in either the buds or the leaves of P. tremuloides (data not shown). The trees at the Mountain Research Station were sampled over the course of the following winter (1991-1992) to determine if acetone was produced only just prior to budbreak or whenever buds were present. While the current year needles continued to produce little or no acetone into the fall, the newly produced buds produced acetone at substantially higher headspace concentrations than were measured from buds during the preceding spring (Fig. 1). Acetone concentrations in these
In this paper, we have presented data that conclusively demonstrate acetone emission by the vegetative tissues of coniferous trees, as measured from both intact and excised tissues. These data appear to be the first documented measurements of the rate of acetone emission from intact tissues. We have also demonstrated that acetone production is seasonal, being emitted by leaves primarily in their primordial state in buds. This result, unfortunately, cannot be compared to the summary results presented by Isidorov et al. [7], as they did not report emission rates nor whether any vegetative buds were present on their branch samples. Acetone emission by buds but not by leaves was independent of the differences in the temperature regimes to which these organs are normally exposed, as this difference was also observed in an isothermal growth chamber experiment. Since acetone is emitted only in trace quantities from conifer leaves (which make up the bulk of the vegetative biomass), it is doubtful that conifer acetone emission contributes significantly to ambient concentrations of acetone in the atmosphere. In animals the production of acetone has been classically associated with the metabolism of acetoacetate derived from excess acetyl-CoA produced during fatty acid degradation [12]. Although a similar metabolic process has not been documented in plants, trees are known to store carbon in the form of lipid [13~-173. In the growth chamber experiment, acetoacetate concentrations
Acetone emission from conifer buds declined after budbreak in a parallel fashion to acetone emission. If acetone is derived from acetoacetate, its synthesis could be enzymatically catalysed by acetoacetate decarboxylase or could result from the spontaneous decarboxylation of acetoacctate [18]. The spontaneous decarboxylation of acetoacetate exhibits a Qio of approximately 5 [18, verified by us]. We measured a QiO for acetone emission from buds of approximately 2, thus indicating that acetone production in the vegetative buds of conifers probably occurs enzymatically. However, we have not been successful thus far in extracting an acetoa&ate decarboxylase activity from coniferous buds. The production of acetone from buds sampled throughout the winter was surprising. Theoretically, if acetone was being produced as a by-product of the degradation of stored lipid, acetone concentrations should be greatest at or around budbreak, when temperatures are warmer and the buds are preparing for leafout. Lipid concentrations are typically highest in midwinter and decline in a parallel fashion with cold hardiness with the onset of spring [19, 203. Instead, acetone concentrations actually declined quite some time prior to budbreak, after a midwinter peak, suggesting that acetone production may have something to do with cold tolerance. However, the mechanism, if any, by which low concentrations of acetone might confer cold tolerance is unknown. These results form the first detailed study of acetone emission from vegetative tissues. While acetone emission from the plants studied here is not sufficient to significantly affect atmospheric chemistry, the pattern of acetone emission raises some intriguing questions as to its role in plant physiology. Though the seasonal pattern of acetone production is suggestive of a role in tolerance to cold temperatures, it is not beyond reason that acetone production may instead be involved somehow in metabolism associated with dormant or meristematic tissues. In this vein, it is noteworthy that the only other documented source of acetone in plants is from seed tissue, another meristematic tissue.
EXPERIMENTAL Species survey. Leaves and leaf buds were sampled from various tree species growing on the University of Colorado, Boulder campus in late May and early June, 1991. This period of time allowed sampling of buds just prior to budbreak and leaves just after budbreak. In addition, vegetative tissues of Populus tremuloides were sampled from greenhouse-grown plants. Excised leaves and buds were incubated at room temp. in 5 ml glass vials (30 ml vials were used with large samples such as needles) sealed with Teflon-lined septa. At least 3 buds were included in each sample vial. After 2 hr (a length of time determined empirically as that required for reaching equilibrium of Me,CO in the headspace of the vial), the headspace gases in the vials were sampled using gas-tight syringes. Headspace gas measurements depend on equilibrium concn relationships between the gas and the liquid phase of the
993
sample at a specific temp. [21]. Therefore, tissue mass is not an important variable in sampling, given that the samples are allowed sufficient time to achieve equilibrium. The time for vegetative buds and leaf samples to achieve equilibrium was determined empirically. Gas chromatography. Headspace gas contents were analysed by GC-FID using a J & W DB-5 0.32mm x 30 m column (WCOT, 1 pm film) at 30” with H, carrier flow of 67 cmsec-‘, or a J &W DB-WAX 0.32 mm x 30 m column (WCOT, 0.25 w film) at 50” with H, carrier at 42 cm set- ‘. Gas samples were cryogenically preconcd in an evacuated Teflon capillary sample loop which was flash heated to 80” prior to loading the sample on to the column. Detector temp. was 200”. The system was calibrated using authentic Me&O standards and the identity of Me&O was confirmed by GC-MS. Surface sterilization. Leaf buds from A. concolor were surface sterilized using 0.1% Triton X-100 and 1% sodium hypochlorite, as described in Kimmerer and MacDonald [21]. After incubating for 2 hr in sealed 30 ml glass vials, headspace Me&O concns were determined as described above. Determination of QIo. To determine the Qi,, of Me&O emission, a modification of the cryotrapping system described by Monson and Fall [22] was used. Excised candles from Pinus Pexilis James were placed in 30 ml glass vials, which were sealed with Teflon-lined septa and maintained at 25” or 35” in a water bath. Charcoalfiltered air was passed through the vials at a flow rate of lOmlmin_ ’ for at least 15 min prior to sampling. During the 6-min sampling period, 50% of the air exiting the vials was passed through a Teflon capillary sample loop submerged in liquid N, and into an evacuated stainless steel flask. At the end of 6 min, the sample loop was flash heated to 80” and the sample was loaded on to the GC system described above. Growth chamber experiment. One-year-old A. concolor seedlings were placed in a growth chamber at 4” day and 8” night temps and a 10 hr photoperiod in order to meet their dormancy requirements. After 4 months, both day and night temps were raised to 25” and the photoperiod lengthened to 18 hr. Me&O emission was measured using a specially constructed acrylic cuvette placed around buds or newly produced needles. Acrylic was previously tested and found to be inert to low concns of gaseous Me&O. Tank air was passed through the cuvette at a flow rate of 80 ml min- ’ and the exiting sample was cryogenically coned and analysed for Me&O by GCFID. The concn of acetoacetate was determined by a modification of the method of Hradecky et al. [23]. Buds were freeze-clamped after determination of Me,CO emission rates, excised, weighed and ground in a mortar and pestle in liquid N,. The powder obtained was then brought up in 5 : 1 (v/w) ice-cold 100 mM CAPS buffer at pH 11. Using a microsyringe, 100 fi of sample was injected through a Teflon-faced septa into each of two 15 ml sample vials containing ca 0.5 g of KzHP04 (5 M, 100 ~1) HsP04 was then injected into one of the vials to catalyse the formation of Me&O from acetoacetate, while 100 A of 100 mM CAPS pH 11, was injected into
994
R. C. MACDONALD and R. FALL
the other vial to determine the background level of Me,CO. Both vials were then incubated at 30” for 4 hr prior to analysis of the headspace gases by CC-FID. The method was calibrated by use of authentic acetoacetate standards. The use of 30” as an incubation temp. was found to avoid the problems caused by degradation of sugars to Me,CO at the higher temps used by Hradecky et al. [23], though longer incubation times were required. Field measurements. Field measurements of Me,CO emissions were made at the University of Colorado Mountain Research Station on trees growing at an elevation of approximately 3000 m (roughly 300 m below treeline). Vegetative tissues of Abies Zusiocarpa, Picea engelmannii, Pinus contorta and P. tremuloides were excised and placed in 5 or 30 ml glass vials. These samples were then brought to the laboratory in Boulder where the headspace gases were analysed by GC-FID. Mean daily air temps were measured at a nearby site by the Mountain Research Station. Acknowledgements-We
thank the University of Colorado Mountain Research Station for providing a sampling site and logistical support, W. Ingino for technical assistance, and P. Goldan for use of his unpublished data and useful discussions. Supported by a Cooperative Institute for Research in Environmental Sciences Visiting Fellowship, and grants ATM-9007849 and ATM-9206621 from the National Science Foundation.
REFERENCES 1. MacDonald, R. C., Kimmerer, T. W. and Razzaghi, M. (1989) Environ. Poll. 62, 337. 2. Tingey, D. T., Turner, D. P. and Weber, J. A. (1991) Trace Gas Emissions by Plants (Sharkey, T. D., Holland, E. A. and Mooney, H. A., eds), pp. 93. Academic Press, San Diego.
Kennedy, R. A., Rumpho, M. E. and Fox, T. C. (1992) Plant Physiol. 100, 1.
Goldan, P., Kuster, W., Montzka, S. and Fehsenfeld, F. (1990) EOS, Trans. Am. Geophys. Union 71, 1228. Fehsenfeld, F., Calvert, J., Fall, R., Goldan, P., Guenther, A. B., Hewitt, C. N., Lamb, B., Liu, S., Trainer, M., Westberg, H. and Zimmerman, P. (1992) Global Biogeochem. Cycles 6, 389.
6. Khalil, M. A. K. and Rasmussen, R. A. (1992) J. Air Waste Manag. Assoc. 42, 810.
7. Isidorov, V. A., Zenkevich, 1. G. and Ioffe, B. V. (1985) Atmos. Environ. 19, 1.
8. Helm, R. E. (1972) Plant Physiol. SO, 293. 9. Fisher, G. S., Legendre, M. G., Lovgren, N. V., Schuller, W. H. and Wells, J. A. (1979) J. Agric. Food Chem. 27, 7.
10. Taylorson, R. B. (1979) Seed Sci. Technol. 7, 369. 11. Murphy, J. B. (1985) Physiol. Plant. 63, 231. 12. Lehninger, A. L. (1982) Principles of Biochemistry. Worth Publishers, New York. 13. Sinnott, E. W. (1918) Botan. Gaz. 66, 162. 14. Robards, A. W. and Kidwai, P. A. (1969) PIanta 84, 239.
Murmanis, L. (1971) Ann. Bot. 35, 133. Hiill, W. (1975) Z. Pjianzenphysiol. 75, 158. Sennerby-Forsse, L. (1986) Physiol. Plant. 67,529. van Stekelenburg, G. J. and Koorevaar, G. (1972) Clin. Chim. Acta 39, 191. 19. Sakai, A. and Larcher, W. (1987) Frost Survival of Plants, pp. 124- 132. Springer, Berlin. 20. Kozlowski, T. T., Kramer, P. J. and Pallardy, S. G. (1991) The Physiological Ecology of Woody Plants, pp. 204206. Academic Press, San Diego. 21. Kimmerer, T. W. and MacDonald, R. C. (1987) Plant 15. 16. 17. 18.
Physiol. 84, 1204.
22. Monson,
R. K. and Fall, R. (1989) Plant Physiol.
90, 267.
23. Hradecky,
P., Jagos,
J. Chromatog. 146, 327.
P. and
Janak,
J. (1978)
1993
ACETONE
0031~9422/93 s6.00+0.00 0 1993 PerganmnPress Ltd
EMISSION FROM CONIFER ROBERT
Cooperative
Institute
C.
MACDONALD
BUDS
and RAY FALL
for Research in Environmental Sciences and Department of Chemistry and Biochemistry, University of Colorado, Campus Box 216, Boulder, CO 80309-0216, U.S.A. (Received
Key Word Index-Abies tolerance; dormancy.
in revised form 5 April 1993)
concolor; Pinaceae; gymnosperm;
conifer; emission; ketone; acetone; bud; cold
Abstract-Acetone was detected in the headspace of excised vegetative buds of all seven conifers examined. Acetone emission rates of 7.4 + 1.5 pg hr- ’ g- 1fr. wt were measured from intact buds of Abies concolor during a growth chamber experiment. Little or no acetone emission was observed from the leaves of conifers or from any angiosperm vegetative tissue tested. Acetone headspace concentrations from buds of three conifer species were measured over two growing seasons at a forest site. Acetone concentrations were highest in winter and dropped precipitously with the onset of budbreak, suggesting that acetone may play a role in cold tolerance in gymnosperms.
INTRODUCTION
Plants are known to emit a large number of volatile compounds, some of which, such as ethylene, ethanol and monoterpenes, are known to play important physiological roles [I -31. Recently, relatively high concentrations of acetone have been measured in the atmosphere of rural pine forests [4-61. The presence of substantial amounts of acetone in rural air raised the question of whether plants could be a significant source of acetone. Isidorov et al. [7] list acetone as being produced by 22 species of plants, as determined by GC-mass spectrometry. Unfortunately, no detailed data were presented; thus, the quantities of acetone emitted and the source of acetone production within the plants are unclear. We could find no reports on the biology of acetone emission from vegetative tissues. There have been several reports in the literature of acetone production by germinating seeds [S-11], possibly in association with lipid metabolism [l 11. Much of the work with acetone production by seeds involved elevated incubation temperatures [9-l 11, leading to the possibility of artifacts such as the thermal decomposition of acetoacetate to acetone. The purpose of this research was to determine whether plants are capable of emitting sufficient amounts of acetone to account for that detected in rural air. We are also interested in determining the role of acetone emissions in plants, and in determining how acetone is produced.
UESULTS
In order to determine whether vegetative tissues were capable of emitting significant quantities of acetone, leaves and buds from trees on the University of Colorado
campus were sampled in a species survey. Little or no acetone was detected in the leaf emissions of any species sampled (Table 1). However, acetone was emitted by the excised vegetative buds of every gymnosperm species studied in headspace concentrations of up to 25.5 ngml-i (Table l), and as high as 200 ngml-’ in later experiments (Fig. 1). Little or no acetone was emitted from the buds of angiosperms (Table 1). Acetone production was not significantly altered by surface sterilization of buds of Abies concolor (P=O.2578, Wilcoxon signed rank test). Acetone production in excised candles of Pinus Jexilis exhibited a Q10 of 2.17kO.54 (Q,e is the factor by which a reaction rate increases with a 10” increase in temperature). To examine whether intact tissues emit acetone, and to determine the effect of budbreak under isothermal conditions, emission rates from intact buds and needles of A. concolor were measured in a growth chamber experiment. At 25”, acetone emission rates were 7.4 f 1.5 pg hr- ’ g- i fr. wt before budbreak, declining to 1.0 +0.2 pghr- 1g - 1fr. wt by the needles produced from these buds (n = 4). Similarly, the concentration of a putative precursor of acetone, acetoacetate, also declined from 46.4k9.2 pgg-l fr. wt in buds to 0.0k4.3 pgg-i fr. wt in the newly-produced needles. To verify the uniqueness of acetone emission to gymnosperm buds in the field, and to more precisely determine the pattern of acetone emission during budbreak, acetone emission from the buds and leaves of Pinus contorta, Abies lasiocarpa, Picea engelmannii and Populus tremuloides was measured over the course of budbreak at a pristine alpine site at the University of Colorado Mountain Research Station in 1991. Substantial production of acetone by the buds of each gymnosperm species was observed, with production declining precipitously 991
R. C. MACDONALDand R. FALL
992
Table 1. Headspace acetone concentrations from leaves and buds of trees Headspace acetone Leaves
Species
Buds (ngml-‘)
Gymnosperms Abies concolor (Gord. & Glend.) Lindl. ex Hildbr. A. lusiocurpa (Hook) Nutt. Picea glauca (Moench) Voss P. pungens Engelm. P. engelmannii Engelm. Pseudotsuga menziesii (Mirb.) Franc0 Pinus contorta Loud.
0.8&0.4 0.2 f 0.2 nd nd _
22.6 + 6.6 25.5 5 2.4 3.8 +0.2 6.8 +0.7 8.2k2.7 2.0 + 0.9 19.3* 7.5
Angiosperms Acer saccharinum L. A. rubrum L. Quercus rubra L. Tilia americana L. Populus tremuloides Michx.
*Data are meanfs.e.
1.1+0.2 nd nd tr nd
tr tr nd
of headspace acetone concentrations produced by three samples
for each species. tr, Trace amounts present; nd, not detectable.
buds increased during the winter, reaching a midwinter peak before declining to values similar to those obtained during the spring of 1991. DISCUSSION
JJASONDJFMAMJJA 1991
I
1992
Fig. 1. Variation in acetone emission from conifer buds with time. Data are mean and s.e. of three samples per species. Arrows indicate where budbreak occurred for all three species.
upon leafout (Fig. 1, inset). An exception was P.contorta, in which some residual acetone emission occurred during the candle stage of development. Acetone production was not detected in either the buds or the leaves of P. tremuloides (data not shown). The trees at the Mountain Research Station were sampled over the course of the following winter (1991-1992) to determine if acetone was produced only just prior to budbreak or whenever buds were present. While the current year needles continued to produce little or no acetone into the fall, the newly produced buds produced acetone at substantially higher headspace concentrations than were measured from buds during the preceding spring (Fig. 1). Acetone concentrations in these
In this paper, we have presented data that conclusively demonstrate acetone emission by the vegetative tissues of coniferous trees, as measured from both intact and excised tissues. These data appear to be the first documented measurements of the rate of acetone emission from intact tissues. We have also demonstrated that acetone production is seasonal, being emitted by leaves primarily in their primordial state in buds. This result, unfortunately, cannot be compared to the summary results presented by Isidorov et al. [7], as they did not report emission rates nor whether any vegetative buds were present on their branch samples. Acetone emission by buds but not by leaves was independent of the differences in the temperature regimes to which these organs are normally exposed, as this difference was also observed in an isothermal growth chamber experiment. Since acetone is emitted only in trace quantities from conifer leaves (which make up the bulk of the vegetative biomass), it is doubtful that conifer acetone emission contributes significantly to ambient concentrations of acetone in the atmosphere. In animals the production of acetone has been classically associated with the metabolism of acetoacetate derived from excess acetyl-CoA produced during fatty acid degradation [12]. Although a similar metabolic process has not been documented in plants, trees are known to store carbon in the form of lipid [13~-173. In the growth chamber experiment, acetoacetate concentrations
Acetone emission from conifer buds declined after budbreak in a parallel fashion to acetone emission. If acetone is derived from acetoacetate, its synthesis could be enzymatically catalysed by acetoacetate decarboxylase or could result from the spontaneous decarboxylation of acetoacctate [18]. The spontaneous decarboxylation of acetoacetate exhibits a Qio of approximately 5 [18, verified by us]. We measured a QiO for acetone emission from buds of approximately 2, thus indicating that acetone production in the vegetative buds of conifers probably occurs enzymatically. However, we have not been successful thus far in extracting an acetoa&ate decarboxylase activity from coniferous buds. The production of acetone from buds sampled throughout the winter was surprising. Theoretically, if acetone was being produced as a by-product of the degradation of stored lipid, acetone concentrations should be greatest at or around budbreak, when temperatures are warmer and the buds are preparing for leafout. Lipid concentrations are typically highest in midwinter and decline in a parallel fashion with cold hardiness with the onset of spring [19, 203. Instead, acetone concentrations actually declined quite some time prior to budbreak, after a midwinter peak, suggesting that acetone production may have something to do with cold tolerance. However, the mechanism, if any, by which low concentrations of acetone might confer cold tolerance is unknown. These results form the first detailed study of acetone emission from vegetative tissues. While acetone emission from the plants studied here is not sufficient to significantly affect atmospheric chemistry, the pattern of acetone emission raises some intriguing questions as to its role in plant physiology. Though the seasonal pattern of acetone production is suggestive of a role in tolerance to cold temperatures, it is not beyond reason that acetone production may instead be involved somehow in metabolism associated with dormant or meristematic tissues. In this vein, it is noteworthy that the only other documented source of acetone in plants is from seed tissue, another meristematic tissue.
EXPERIMENTAL Species survey. Leaves and leaf buds were sampled from various tree species growing on the University of Colorado, Boulder campus in late May and early June, 1991. This period of time allowed sampling of buds just prior to budbreak and leaves just after budbreak. In addition, vegetative tissues of Populus tremuloides were sampled from greenhouse-grown plants. Excised leaves and buds were incubated at room temp. in 5 ml glass vials (30 ml vials were used with large samples such as needles) sealed with Teflon-lined septa. At least 3 buds were included in each sample vial. After 2 hr (a length of time determined empirically as that required for reaching equilibrium of Me,CO in the headspace of the vial), the headspace gases in the vials were sampled using gas-tight syringes. Headspace gas measurements depend on equilibrium concn relationships between the gas and the liquid phase of the
993
sample at a specific temp. [21]. Therefore, tissue mass is not an important variable in sampling, given that the samples are allowed sufficient time to achieve equilibrium. The time for vegetative buds and leaf samples to achieve equilibrium was determined empirically. Gas chromatography. Headspace gas contents were analysed by GC-FID using a J & W DB-5 0.32mm x 30 m column (WCOT, 1 pm film) at 30” with H, carrier flow of 67 cmsec-‘, or a J &W DB-WAX 0.32 mm x 30 m column (WCOT, 0.25 w film) at 50” with H, carrier at 42 cm set- ‘. Gas samples were cryogenically preconcd in an evacuated Teflon capillary sample loop which was flash heated to 80” prior to loading the sample on to the column. Detector temp. was 200”. The system was calibrated using authentic Me&O standards and the identity of Me&O was confirmed by GC-MS. Surface sterilization. Leaf buds from A. concolor were surface sterilized using 0.1% Triton X-100 and 1% sodium hypochlorite, as described in Kimmerer and MacDonald [21]. After incubating for 2 hr in sealed 30 ml glass vials, headspace Me&O concns were determined as described above. Determination of QIo. To determine the Qi,, of Me&O emission, a modification of the cryotrapping system described by Monson and Fall [22] was used. Excised candles from Pinus Pexilis James were placed in 30 ml glass vials, which were sealed with Teflon-lined septa and maintained at 25” or 35” in a water bath. Charcoalfiltered air was passed through the vials at a flow rate of lOmlmin_ ’ for at least 15 min prior to sampling. During the 6-min sampling period, 50% of the air exiting the vials was passed through a Teflon capillary sample loop submerged in liquid N, and into an evacuated stainless steel flask. At the end of 6 min, the sample loop was flash heated to 80” and the sample was loaded on to the GC system described above. Growth chamber experiment. One-year-old A. concolor seedlings were placed in a growth chamber at 4” day and 8” night temps and a 10 hr photoperiod in order to meet their dormancy requirements. After 4 months, both day and night temps were raised to 25” and the photoperiod lengthened to 18 hr. Me&O emission was measured using a specially constructed acrylic cuvette placed around buds or newly produced needles. Acrylic was previously tested and found to be inert to low concns of gaseous Me&O. Tank air was passed through the cuvette at a flow rate of 80 ml min- ’ and the exiting sample was cryogenically coned and analysed for Me&O by GCFID. The concn of acetoacetate was determined by a modification of the method of Hradecky et al. [23]. Buds were freeze-clamped after determination of Me,CO emission rates, excised, weighed and ground in a mortar and pestle in liquid N,. The powder obtained was then brought up in 5 : 1 (v/w) ice-cold 100 mM CAPS buffer at pH 11. Using a microsyringe, 100 fi of sample was injected through a Teflon-faced septa into each of two 15 ml sample vials containing ca 0.5 g of KzHP04 (5 M, 100 ~1) HsP04 was then injected into one of the vials to catalyse the formation of Me&O from acetoacetate, while 100 A of 100 mM CAPS pH 11, was injected into
994
R. C. MACDONALD and R. FALL
the other vial to determine the background level of Me,CO. Both vials were then incubated at 30” for 4 hr prior to analysis of the headspace gases by CC-FID. The method was calibrated by use of authentic acetoacetate standards. The use of 30” as an incubation temp. was found to avoid the problems caused by degradation of sugars to Me,CO at the higher temps used by Hradecky et al. [23], though longer incubation times were required. Field measurements. Field measurements of Me,CO emissions were made at the University of Colorado Mountain Research Station on trees growing at an elevation of approximately 3000 m (roughly 300 m below treeline). Vegetative tissues of Abies Zusiocarpa, Picea engelmannii, Pinus contorta and P. tremuloides were excised and placed in 5 or 30 ml glass vials. These samples were then brought to the laboratory in Boulder where the headspace gases were analysed by GC-FID. Mean daily air temps were measured at a nearby site by the Mountain Research Station. Acknowledgements-We
thank the University of Colorado Mountain Research Station for providing a sampling site and logistical support, W. Ingino for technical assistance, and P. Goldan for use of his unpublished data and useful discussions. Supported by a Cooperative Institute for Research in Environmental Sciences Visiting Fellowship, and grants ATM-9007849 and ATM-9206621 from the National Science Foundation.
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