Environ. Sci. Technol. 2005, 39, 268-273
Real-Time Visualization and Quantification of PAH Photodegradation on and within Plant Leaves EDWARD WILD, JOHN DENT, GARETH O. THOMAS, AND KEVIN C. JONES* Departments of Environmental and Biological Sciences, Lancaster University, Lancaster LA1 4YQ, U.K.
Vegetation plays a key role in the environmental cycling and fate of many organic chemicals. A compound’s location on or within leaves will affect its persistence and significance; retention in surface compartments (i.e., the epicuticular wax and cuticle) renders the compound more susceptible to photodegradation and volatilization, while penetration into the epidermal cell walls or cytoplasm will enhance susceptibility to metabolism. Here, for the first time, methodologies which combine plant and PAH autofluorescence with two-photon excitation microscopy (TPEM) are used to visualize and quantify compound photodegradation on and within living plant leaves. Anthracene, fluoranthene, and phenanthrene were introduced to living leaves of Zea mays and monitored in real time, in control treatments, and when subject to UV-A radiation. Compound photodegradation was observed directly; different degradation rates occurred for different compounds (anthracene > fluoranthene > phenanthrene) and in different locations (at the leaf surface > within the epidermal cells). Results suggest that photodegradation on vegetation may be a more important loss mechanism for PAHs than previously thought. Compound fate in vegetation is potentially highly complex, influenced by diffusion into and location within leaf structures, the rates of supply/loss with the atmosphere, exposure to sunlight, and other environmental conditions. The techniques described here provide a real-time tool to advance insight into these issues.
Introduction Over 80% of the earth’s terrestrial surface is covered by vegetation (1). The large leaf area of many species/ecosystems provides an extensive surface for gaseous exchange with the atmosphere, and exposure to sunlight (1, 2). Leaves are coated by a hydrophobic lipid layer, the cuticle (2), which can accumulate many organic compounds from the atmosphere, or agrochemicals added to the plant surface (3-5). Compounds may be stored there, migrate to the inner leaf, undergo degradation (photolytic; metabolic), or re-release to the atmosphere (6-8). The relative importance of these processes is controlled by the location of the compound (in turn controlled by properties of the compound and the plant), and environmental conditions. For example, * Corresponding author phone: (44)1524-593972; fax: (44)1524593985; e-mail:
[email protected]. 268
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retention in surface compartments (i.e., the epicuticular wax and cuticle) may render the compound more susceptible to photodegradation and volatilization, while penetration to the epidermal cell walls or cytoplasm will enhance susceptibility to metabolism, and any potentially adverse effects of the compound. Vegetation-compound interactions are therefore pivotal to the global fate, behavior, and significance of many organic chemicals, including persistent organic pollutants (POPs) (9, 10). POPs become sorbed to the leaf surface at the epicuticular wax, the outermost region of the cuticle. The plant cuticle is a hydrophobic lipid structure, typically 0.1-10 µm thick, synthesized by the epidermal cells (11, 12). POPs would need to diffuse through this and the cuticular matrix (comprising the epicuticular wax, the cuticle proper, the cuticle layer, the pectinous layer, and the cell wall) (12) before reaching the cytoplasm of the epidermal cells. The light dependence of plants means their leaves can be subject to sustained solar light intensities. This, combined with the large surface area of vegetative cover noted above, provides the scope for compound photolysis. The solar spectrum at the earth’s surface constitutes UV-B radiation (280-320 nm), of which ∼99% is removed in the stratosphere, UV-A radiation (320-400 nm), of which ∼6% reaches the earth’s surface, and visible light at 400-785 nm. Photolysis reactions represent an important transformation pathway for some POPs, notably many polynuclear aromatic hydrocarbons (PAHs) (7, 8). PAH photodegradation is known to be highly dependent upon substrate. Fly ash and carbon black may “protect” PAHs from photolysis, for example (12-14). Faster photolysis has been reported for PAHs on spruce needles than on fly ash or carbon black (7, 8). Plant tissue is an optically complex medium (15), likely to influence the rates of photodegradation through modification of solar radiation to separate desirable wavelengths from those which are detrimental to the plants health. The plant cuticle filters, scatters, and focuses certain wavelengths (2). The epicuticular wax and cuticle induce intense reflection and refraction of the visible wavelengths (particularly when high proportions of crystalline wax are present) and absorb UV radiation (15-17). UV radiation is attenuated within the cuticle and epidermal cells more strongly than visible radiation (16), with up to 55% of UV radiation being attenuated within the first few micrometers of the leaf (2, 17). However, some UV-A is not filtered by the outermost layers of a leaf and can still penetrate deep into internal structures, up to 250 µm in fir and 450 µm in spruce (17). UV radiation can itself be detrimental to plant growth, by reducing chlorophyll and photosystem two activity (2); it can also induce leaf and cuticular wax thickening, and epidermal cell and DNA damage (2, 18). We have previously described how TPEM techniques can help studies to visualize and track organic compounds in living plants using plant and compound autofluorescence (19). TPEM is a new form of laser scanning microscope that enables identification of chemical and structural components within a sample, through the utilization of their fluorescence excitation and emission spectra (20). In our previous study (19), the PAH anthracene was observed within the epicuticular wax, cuticle, epidermal cell walls, and cytoplasm of living leaves and diffusion rates derived for its movement through the surface layers. Here, we further demonstrate the tremendous scope of this technique by monitoring the realtime photodegradation of three PAHs, anthracene, fluoranthene, and phenanthrene, at the leaf surface and within the epidermal cells of living maize (Zea mays) leaves when 10.1021/es0494196 CCC: $30.25
2005 American Chemical Society Published on Web 11/24/2004
irradiated with UV-A radiation. TPEM provides a method to image living cells and produce three-dimensional images with little risk of photobleaching or phototoxicity to the sample. Two-photon excitation of fluorescent molecules is related to the simultaneous (10-16 s) absorption of two low-energy photons, which together provide the higher energy required to induce fluorescence (20). The two photons can only combine at the focal plane, to induce excitation, if they interact simultaneously. Two-photon excitation has the important consequence of limiting the excitation region within the sample to within a subfemtoliter volume, thereby eliminating background fluorescence, increasing sample penetration, and allowing 3D imaging. Using TPEM, sample fluorescence can be achieved at a shorter wavelength than the excitation wavelength, where, for instance, two 700 nm photons can combine to induce excitation at 350 nm and fluorescence at 400-500 nm. Bleaching of the sample is reduced due to the use of an infrared laser, allowing increased periods of analysis. 3D sectioning can be achieved through the successive adjustment of the focal plain within a sample making a series of successive images through the sample This can be done in both xy and xz planes. A 3D reconstruction of the sample can then be produced using the appropriate computing/software techniques (20).
Materials and Methods Experimental Section. Plants. Seeds of Zea mays cultivar gl 26 were obtained from the Maize Genetics Cooperation Stock Center, IL. All seeds were backcrossed to the same line for 5 generations, for genetic homogeneity. Plants were grown in Levington compost (original compost mix, M3), under a 16 h photoperiod, illuminated by 400 W Na solar lighting. Plants were watered four times a day for 5 min periods from below using a capillary matting dripper system. The greenhouse temperature was maintained at ∼25 °C. Plants 21-25 days old were used for the experiments. Compounds and Solution Generation. Anthracene, fluoranthene, and phenanthrene were obtained from Aldrich Chemical Co. at 99.9%, 98%, and 99.9% purity, respectively. First, 1 mg of each compound was placed in a separate 10 mL vial with 10 mL of acetone. The vial was placed in an amber container, to reduce any risk of photodegradation. This was placed in a Branson 3210 sonic bath for 45 min at an RF frequency of 47 kHz ( 6% to aid solubilization. Preparation of Control Slides. Compounds were applied in solution as a homogeneous layer to solvent-rinsed BDH super premium microscope slides using a micropipet, typically at ∼0.3 µg cm-2. These were “controls” to ascertain compound photodegradation rates independently from when they were added to the plant leaves. The slides were treated identically to the plants prior to analysis. Plant Contamination. Regions ∼16 cm2 were selected for contamination on leaves 4-6 of 21-25 day old plants. Compounds were applied in solution as a homogeneous layer to the underside of separate leaves of living plants, as described above. The compound was applied to the underside of the leaf, to ensure they could be attached to the microscope without damaging the plant. The upper surface of the leaf was attached to a microscope slide; this kept the sample flat, enabling the lower surface of the leaf to be in direct contact with the water immersion lens, without needing to bend or damage the living leaf. The micropipet did not come into direct contact with the leaf, to avoid abrasion. Postcontamination, the plants were maintained in an indoor ambient environment at 25 °C under diffuse Na solar lighting, prior to analysis. The contaminated plants were not exposed to direct solar radiation. Living plants were analyzed either 0.5 h postcontamination to study the compound at the leaf surface, or after 72 h to study the compound within the cell
walls or cytoplasm of the epidermal cells, by which time they had reached a depth of ∼22-26 µm. It had previously been established that this time allowed the compound to penetrate to these depths in the leaves (19). Instrumentation. A Bio-Rad Radiance 2000 MP scanning system was used with a Spectra Physics Tsunami/millennia laser 690-1050 nm and a Nikon Eclipse TE300 inverted microscope. The laser wavelength was set to 700 nm. Images were collected and processed using Bio-Rad Lasersharp 2000 imaging software and Confocal assistant 4.02. Fluorophore-containing compounds fluoresce when excited by a range of wavelengths. Specific wavelengths can be selected to excite a number of different fluorophores simultaneously, which individually will have unique emission spectra, allowing their unambiguous identification. Anthracene, fluoranthene, and phenanthrene were independently identified in living leaves of maize using fluorescence. Plantspecific autofluorescence excitation and emission profiles were determined when excited at 350 nm using TPEM, allowing the visualization of the compounds of interest within the leaf. Emission detection filters were set to detect anthracene and phenanthrene at 390 ( 35 nm (HQ390/70), using blue as a pseudo color channel, and fluoranthene, using blue/green as a pseudo color channel. Cell walls, the cuticle, stomata, hairs, and cellular vacuoles were detected at 528 ( 25 nm (HQ528/50), with chloroplasts and organelles detected at 590 ( 35 nm (HQ590/70) using green and red as pseudo color channels, respectively. All studies were performed with a Nikon ×20/0.75 Plan Fluor D.I.C. water immersion lens. Irradiation Experiments. TPEM was used to study the photodegradation of anthracene, phenanthrene, and fluoranthene, when subjected to irradiation with a 100 W Nikon super high-pressure mercury lamp, passed through both a Nikon 4 and 8 medium-density filter and a Nikon UV-1A dichroic mirror DM400 Ex 365/10 BA 400, to control both the UV intensity and the irradiance wavelength spectra, allowing 355-375 nm excitation wavelengths to pass through. Timecourse experiments were performed with detection in real time. Each sample was exposed to 45 × 60 s cycles of UV-A radiation, with 30 s of irradiation on and 30 s of irradiation off in every cycle. The time off was used to scan the image. Photodegradation rates were determined for compounds on the slides and on/in the leaves at ∼0-3 µm from the leaf underside surface and within the epidermal cells at a depth of ∼22-26 µm. The studies were performed for each compound at least three times on separate leaves in separate time-course experiments, at both the leaf surface and within the epidermal cells. For each experiment, 6-9 “regions of interest” (ROI) were selected and monitored in each sample. These encompassed the compound on or within the leaf and uncontaminated areas, to provide both compound degradation data and leaf background data during the same experimental run. This approach is illustrated in Figure 1. The photodegradation rate was determined through the use of mean pixel intensity within the ROI, using 256 different intensities per pixel. The ROI were selected to either encompass areas of PAHs to study degradation, or areas devoid of PAHs (control) to study the plants background fluorescence. Control experimental runs were performed without UV-A radiation, for both contaminated and uncontaminated plants (“dark runs”), to determine plant fluorescence degradation or loss of compound induced by the laser. The laser induced no degradation of the compound. Uncontaminated control plants were studied to determine if the plant autofluorescence decreased under UV-A radiation. The plants autofluorescence was unmodified during the experimental period. The laser was set to 20% power (as the output power of the laser) to analyze anthracene and fluoranthene, and 30% VOL. 39, NO. 1, 2005 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 1. Autofluorescence of the surface of a leaf of Zea mays contaminated with anthracene. XY cross-sectional image showing anthracene (blue) bound at the leaf surface using a scanned width of 416 µm, with a ×20/0.75 plan fluor D.I.C. water imersion lense and zoom factor of 1.5. RGB color mode filters: red HQ 590 ( 35 nm; green HQ 528 ( 25 nm; blue HQ 390 ( 35 nm, excited using two-photon excitation at 700 nm. Anthracene appears as blue, with the cuticle and cell walls shown green and chloroplasts within the stomata shown as yellow/red. Typical regions of interest (ROI) selected for study are shown. ROI 1-5 represent regions of anthracene at the leaf surface selected for photodegradation quantification. ROI 6-8 represent uncontaminated areas on the same leaf to show leaf background fluorescence data compound in relation to compound degradation data. for phenanthrene A; higher laser power was needed to image phenanthrene as its fluorescence signal was weaker than the other two compounds. XZ images were made before and after each study to determine if the sample moved during the analysis period. If leaf movement was perceived, the sample was discarded. The UV intensities were recorded, using a double monochromator SR9910 spectroradiometer (Macam Photometrics, Scotland) between 350 and 380 nm with a highest intensity of 24 W m-2. The UV-A intensities were recorded at the front focal plane of the lens to be as close to the illumination received by the sample as possible. The UV-A light intensities used ensured that photodegradation could be quantified and visualized over a short time period and that optical filtering and photodegradation rates between the leaf surface and epidermal cells of the leaf could be accurately studied. This was limited by the available light source and period over which the living plants could be analyzed. Statistical Analysis. Photodegradation was first order; loss rates and half-lives were determined using SPSS 11.5.e. Independent T-test analysis was carried out for data determined at the leaf surface and internally, to ascertain if the values were significantly different. Residuals analysis was 270
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performed and did not indicate significant deviation from the first-order assumption.
Results and Discussion Introductory Remarks. Anthracene, fluoranthene, and phenanthrene exhibited pseudo first-order degradation when subject to UV-A irradiation. This was observed on the glass slides, and at the surface and interior of the leaves. Figure 2 shows illustrative images of the changing amounts of the three compounds detected in the surface 0-3 µm of leaves during typical 45 min time-course experiments. Figure 3 shows images through the leaf (XZ images), to highlight the occurrence of anthracene at the surface before and after exposure to UV-A radiation. Experimental runs, without UV-A radiation (“dark runs”), showed no degradation or loss of compound with compound fluorescence intensities remaining uniform throughout the run period. Compound volatilization from the leaf surface could be ruled out, because the leaves were in direct contact with a layer of water, separating the sample and the water immersion lens. Detectable quantities of compound did not move into the water layer; this was analyzed through wide field fluorescence microscopy, using the 100 W Nikon super high-pressure mercury
FIGURE 2. Montage of images showing the degradation of anthracene (a), fluoranthene (b), and phenanthrene (c) at the leaf surface over 45 min. 3 × 6 XY cross-sectional images are shown, taken over a 45 min time-course study when subjected to UV-A radiation. Anthracene (a) is shown as blue and can be seen to rapidly reduce in intensity over time. Fluoranthene (b) is shown as a diffuse area of blue/green clusters which reduce in intensity from t ) 0 to t ) 2700. Phenanthrene (c) is shown as blue; little change in intensity is visible for phenanthrene. The chloroplast brightness in image a intensifies throughout the time-course experiment due to photolytic activation induced by the UV-A radiation.
FIGURE 3. Cross-sectional (XZ) images showing anthracene at the leaf surface, before (a) and after (b) photodegradation using UV-A radiation. These images correspond to the section shown in Figure 2. XZ cross-sectional images through Figure 2, shown using a scanned width of 416 µm and a depth of 73 µm, with a step of 1.5 µm and a zoom factor of 1.5. Colors are as noted for Figure 1. UV lamp. Any fluorescence could be rapidly identified. The way in which the compounds degraded appeared similar, with the compound fluorescence steadily dimming before “disappearing”. Derivation of Half-Lives and Comparison of Surface and Subsurface Degradation Rates. Photodegradation rates on the glass slides averaged (( standard deviation) 2.2 ( 0.6, 5.9 ( 0.1, and 22.6 ( 4.3 min for anthracene, fluoranthene, and phenanthrene, respectively. Values determined on leaf surfaces (0-3 µm) closely resembled these, 2.9 ( 0.4, 5.1 ( 0.7, and 27.4 ( 8.1 min for anthracene, fluoranthene, and phenanthrene, respectively. Photodegradation rates determined in the epidermal cells were significantly slower (at the 99% level) for anthracene and fluoranthene, by factors of 1.5 and 2.7, respectively. Figure 4 shows some illustrative timecourse data. This is likely due to modifications of the optical properties and intensity of the radiation reaching the subsurface layers (15-17). As noted earlier, UV radiation is attenuated within the cuticle and epidermal cells more strongly than visible radiation (15, 17) with up to 55% reduced within a few micrometers of the leaf surface (2, 17). A
proportion of UV-A not attenuated by the outermost tissues of a leaf can still penetrate deep into its internal structure up to 250 µm in fur and 450 µm in spruce (17). Phenanthrene half-lives could not be determined within the epidermal cells because it persisted longer than the duration of the experiment. Comment on the Compound Differences. Table 1 presents a summary of measurements made here on maize and other data determined under different experimental conditions on spruce needles (7, 8). It is stressed that the absolute half-life values determined in this and other investigations will be a function of the irradiation intensity and other environmental conditions. Some studies do not appear to adequately control for other losses. It is therefore unwise to compare these directly across studies. However, the order of reactivities observed here, anthracene > fluoranthene > phenanthrene, clearly mirrors that seen elsewhere (7, 8, 21). Comments on the Rates and Relationship to Field Conditions. The UV-A intensities used in this study were high, as mentioned earlier. Typical values for temperate VOL. 39, NO. 1, 2005 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 4. Typical photodegradation data generated during the study.
TABLE 1. Photodegradation Rates Determined in This Study, Normalized to Average Conditions and Related to Other Studies photodegradation rates half-lives (h) normalized dataa
measured
literature data for spruce needles
compound
surface
cellular
surface
cellular
ref 7
ref 8
anthracene fluoranthene phenanthrene
0.05 ( 0.00076 0.09 ( 0.00038 0.46 ( 0.00015
0.08 ( 0.00085 0.24 ( 0.00023
5.4 9.6 52
9.2 27
25 26 75
170 180 230
a Normalized for temperature and light, to derive photodegradation half-lives on leaves representative of average northern hemisphere conditions (see text for details).
environmental conditions average a few W m-2 in temperate latitudes and may reach 10 or more W m-2 under the most intense tropical conditions (22). The photodegradation rates measured here were therefore rapid, akin to values that have been reported previously for the photodegradation of PAHs in water (21, 23). If it is assumed that the relationship between the rate of photodegradation and light intensity is linear, and the data are “normalized” to represent typical solar radiation values averaged for the northern hemisphere (22), the photodegradation rates measured here are within a factor of 2-9 of those measured by others on vegetation (7, 8). The experiments conducted here were performed at 25 °C, which may give an approximate 2-fold increase over the rate reported for average northern hemisphere conditions. Thus, normalizing the data to light intensity and temperature gives rates of degradation within a factor of 1-5 of those measured in other vegetation studies (see Table 1) (7, 8). Comments on the Implications for the Presence/ Processing of PAHs in the Environment. PAHs reach leaves primarily from the atmosphere, from where they are supplied in gaseous and particulate forms (1, 3, 24). It is generally assumed that leaf concentrations approach equilibrium with gas-phase concentrations over time (25, 26), although uptake may be kinetically constrained during the lifetime of leaves/needles by the rates of supply from the atmosphere (25, 27). However, in practice, we have observed that PAHs and a range of other POPs may approach “steadystate” leaf concentrations over a few days (28, 29). The data from this study show that PAH photodegradation is a potentially rapid loss process which is likely to have an important influence over leaf concentrations in the field. Given the rate at which photodegradation can occur, it is pertinent to consider how/why PAHs continue to be present in leaves. Presumably, the answer is because (a) they are 272
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constantly generated and supplied fresh from the atmosphere; (b) it may be dark/cloudy, when they are supplied to the leaf; (c) once they start to penetrate into the leaf, they become more protected from photodegradation; (d) many leaves are “self-shaded” in the field or are receiving only a tiny fraction of the irradiances applied here; and (e) PAHs also arrive on particles, and may be protected while residing upon them (13, 14). In summary, these data indicate that photodegradation on vegetation may be important to the global cycling fate of POPs. However, it will be complicated to include this loss pathway in mass balances/environmental inventories because the rate of loss presumably depends on light intensities in the field, species and habitat differences, compound isolation within the leaves, and the properties of individual POP compounds. The ability to noninvasively visualize the degradation of a compound on or within a living plant has enabled the first direct quantitative determination of this loss process. The results presented provide an initial insight into the complexities involved in compound photodegradation when it is adhered to a plant. Limitations to the Technique and Future Work. The technique currently has a number of limitations; it is not as sensitive as some current analytical procedures for detecting POPs, and it relies specifically on chemicals/ compounds which fluoresce, meaning certain compounds cannot be analyzed. However, it opens up a new visual perspective to the study of organic pollutants within the living matrix of plants, helping to gain an understanding of exactly how compounds behave within plants, and where they are located. Through continued development, this technique will provide much valuable information regarding the processing of organic pollutants by plants and other matrixes.
Acknowledgments We are grateful to the U.K. Natural Environment Research Council (NERC) for provision of a Ph.D. studentship (NER/ S/A2002/10394) to E.W.
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Literature Cited (1) Simonich, S. L.; Hites, R. A. Vegetation-atmosphere partitioning of polycyclic aromatic hydrocarbons. Environ. Sci. Technol. 1994, 28, 939-943. (2) Kerstiens, G. The Plant Cuticle: An Integrated Functional Approach; Bios Scientific Publishers: Oxford, 1996. (3) Simonich, S. L.; Hites, R. A. Importance of vegetation in removing polycyclic aromatic hydrocarbons from the atmosphere. Nature 1994, 370, 49-51. (4) Wania, F.; Mackay, D. Tracking the distribution of persistent organic pollutants. Environ. Sci. Technol. 1996, 30, 391A-396A. (5) Kirkwood, R. C. In Pesticides on Plant Surfaces; Cottrell, H. J., Ed.; John Wiley and Sons: New York, 1987; pp 1-25. (6) Smith, K. E. C.; Jones, K. C. Particles and vegetation: Implications for the transfer of bound organic contaminants to vegetation. Sci. Total Environ. 2000, 246, particle-207-236. (7) Niu, J.; Chen, J.; Martens, D.; Quan, X.; Yang, F.; Kettrup, A.; Schramm, K.-W. Photolysis of polycyclic aromatic hydrocarbons adsorbed on spruce [Picea abies (L.) Karst.] needles under sunlight. Environ. Pollut. 2003, 123, 39-45. (8) Niu, J.; Chen, J.; Martens, D.; Henkelmann, B.; Quan, X.; Yang, F.; Seidlitz, H. K.; Schramm, K.-W. The role of UV-B on the degradation of PCDD/Fs and PAHs sorbed on surfaces of spruce (Picea abies (L.) Karst.) needles. Sci. Total Environ. 2004, 322, 231-241. (9) Wania, F.; McLachlan, M. Estimating the influence of forests on the overall fate of semivolatile organic compounds using a multimedia fate model. Environ. Sci. Technol. 2001, 35, 582590. (10) Wania, F.; Mackay, D. Tracking the distribution of persistent organic pollutants. Environ. Sci. Technol. 1996, 30, 391A-396A. (11) Kirkwood, R. C. Recent developments in our understanding of the plant cuticle as a barrier to the foliar uptake of pesticides. Pestic. Sci. 1999, 55, 69-77. (12) Behymer, T. D.; Hites, R. A. Photolysis of polycyclic aromatic hydrocarbons absorbed on simulated atmospheric particulates. Environ. Sci. Technol. 1985, 19, 1004-1006. (13) Korfmacher, W. A.; Natusch, D. F. S.; Taylor, D. R.; Mamantov, G.; Wehry, E. L. Oxidative transformation of polycyclic aromatic hydrocarbons adsorbed on coal fly ash. Science 1980, 207, 763765. (14) Korfmacher, W. A.; Wehry, E. L.; Mamantov, G.; Natusch, D. F. S. Resistance to photochemical decomposition of polycyclic aromatic hydrocarbons vapor-adsorbed on coal fly ash. Environ. Sci. Technol. 1980, 14, 1094-1099. (15) Yan-Ping, C.; Bornman, F. J. The effect of exposure to enhanced UV-B radiation on the penetration of monochromatic and
(18) (19)
(20) (21) (22) (23)
(24) (25) (26)
(27)
(28) (29)
polychromatic UV-B radiation in leaves of Brassica napus. Physiol. Plant. 1993, 87, 249-255. Bornman, F. J.; Vogelmann, T. C. Effect of UV-B radiation on leaf optical properties measured with fiber optics. J. Exp. Bot. 1991, 42, 547-554. Bornman, F. J.; Vogelmann, T. C. Penetration of blue and UV radiation measured by fiber optics in spruce and fir needles. Physiol. Plant. 1988, 72, 699-705. Kakani, V. G.; Reddy, K. R.; Zhao, D.; Sailaja, K. Field crop responses to ultraviolet-B radiation: a review. Agric. For. Meteorol. 2003, 120, 191-218. Wild, E.; Dent, J.; Barber, J. L.; Thomas, G. O.; Jones, K. C. A novel analytical approach for visualizing and tracking organic chemicals in plants. Environ. Sci. Technol. 2004, 38, 41954199. Diaspro, A. Confocal and Two Photon Microscopy Foundations, Applications, and Advances; Wiley-Liss, Inc: New York, 2002. Mackay, D.; Shiu, W. Y.; Ma, K. C. Physical Chemical Properties and Environmental Fate Handbook; Chapman & Hall/ CRCnetBase: New York, 2000. Seckmeyer, G.; Mayer, B.; Erb, R.; Bernhard, G. Geographical differences in the UV measured by intercompared spectroradiometers. Geophys. Res. Lett. 1995, 21, 577-580. Lehto, K.-M.; Vuorimaa, E.; Lemmetyinen, H. Photolysis of polycyclic aromatic hydrocarbons (PAHs) in dilute aqueous solutions detected by fluorescence. J. Photochem. Photobiol., A: Chem. 2000, 136, 53-60. Howsam, M.; Jones, K. C.; Ineson, P. PAHs associated with the leaves of three deciduous tree species. II: Uptake during a growing season. Chemosphere 2001, 44, 155-164. McLachlan, M. S. Framework for the interpretation of measurements of SOCs in plants. Environ. Sci. Technol. 1999, 33, 17991804. Barber, J. L.; Thomas, G. O.; Kerstiens, G.; Jones, K. C. Current issues and uncertainties in the measurement and modeling or air-vegetation exchange and within -plant processing of POPs. Environ. Pollut. 2004, 128, 99-138. Hellstrom, A.; Kylin, H.; Strachan, W. M. J.; Jensen, S. Distribution of some organochlorine compounds in pine needles from Central and Northern Europe. Environ. Pollut. 2004, 128, 29-48. Smith, K. E. C.; Thomas, G. O.; Jones, K. C. Seasonal and species differences in the air-pasture transfer of PAHs. Environ. Sci. Technol. 2001, 35, 2156-2165. Thomas, G.; Sweetman, A. J.; Ockenden, W. A.; Mackay, D.; Jones, K. C. Air-pasture transfere of PCBs. Environ. Sci. Technol. 1998, 32, 936-942.
Received for review April 17, 2004. Revised manuscript received September 30, 2004. Accepted October 12, 2004. ES0494196
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