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Critical Review
Phytovolatilization of Organic Contaminants Matt Alan Limmer, and Joel Gerard Burken Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.5b04113 • Publication Date (Web): 01 Jun 2016 Downloaded from http://pubs.acs.org on June 2, 2016
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Environmental Science & Technology
PHYTOVOLATILIZATION OF ORGANIC CONTAMINANTS
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Matt Limmer1* & Joel Burken2
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1
University of Delaware, Department of Plant & Soil Sciences, Newark, DE
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2
Missouri University of Science and Technology, Department of Civil, Architectural and Environmental
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Engineering, Rolla, MO
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* Corresponding Author:
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[email protected] 419.276.5358
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152 Townsend Hall, Newark, DE 19716
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ABSTRACT
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Plants can interact with a variety of organic compounds, and thereby affect the fate and transport of
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many environmental contaminants. Volatile organic compounds may be volatilized from stems or leaves
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(direct phytovolatilization) or from soil because of plant root activities (indirect phytovolatilization).
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Fluxes of contaminants volatilizing from plants are important across scales ranging from local
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contaminant spills to global fluxes of methane emanating from ecosystems biochemically reducing
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organic carbon. In this article past studies are reviewed to clearly differentiate between direct- and
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indirect-phytovolatilization and discuss the plant physiology driving phytovolatilization in different
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ecosystems. Current measurement techniques are also described, including common difficulties in
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experimental design. We also discuss reports of phytovolatilization in the literature, finding that
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compounds with low octanol-air partitioning coefficients are more likely to be phytovolatilized (log KOA
50 Pa) of subsurface tissues.32-35
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These convective flows are thought to be driven by differences in humidity between the humid plant
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interior and the relatively dry atmosphere. With flows exceeding 1 mL/min during these convective
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events, advective flux of VOCs in these plants may significantly contribute to direct phytovolatilization
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flux.
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In herbaceous plants, during transport of VOCs to aboveground tissues these compounds can
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escape from primary tissues such as stems and leaves. The plant epidermis is covered by a cuticle, which
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consists of hydrophobic waxes and cutin to limit water loss. The cuticle is occasionally interrupted by
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stomata, which open and close to regulate gas exchange. Thus, transport of VOCs through the epidermis
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occurs through two parallel pathways: diffusion through open stomata and diffusion across the cuticle.
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Permeability of the cuticle is enhanced for molecules that strongly partition into the cuticle and have
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small molar volumes.36, 37 Permeability of the cuticle is also strongly plant and temperature
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dependent,36, 37 and various regression models have been developed to predict permeability of plant
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cuticles.37-39
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In woody species, direct phytovolatilization from tree trunks and branches involves the diffusion
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of VOCs through secondary xylem, secondary phloem and periderm, eventually reaching the
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atmosphere where activity of the VOC is assumed lower than in the tree. The outermost layer of the
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periderm, the phellem, consists of dead cells filled with suberin and waxes to regulate water loss from
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the trunk. However, the phellem must allow some exchange of gasses to support the respiration of living
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cells in the trunk. To allow gas exchange, the phellem is interrupted by lenticels, which are more
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permeable to oxygen and water vapor than the surrounding tissue,40 although the permeance of
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lenticels to H2O and O2 has been measured at ~1 order of magnitude lower than stomata.40 Interestingly,
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lenticels appear to open and close seasonally, reducing the trunk’s permeability during the winter.41 A
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study with evergreen species found no seasonal change in lenticel morphology, while deciduous species
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exhibited seasonal changes in lenticel morphology.42 Further complicating gas transport in trees, some
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trees growing in waterlogged conditions develop aerenchymatous phellem (secondary aerenchyma) to
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aid in oxygen transport to the roots.43-45 The prevalence of aerenchymatous phellem and its effect on
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VOC transport in wood remains uncertain; however, increasing the transport of O2 will certainly increase 8
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transport of other volatile molecules. Plants growing in flooded conditions can also develop
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hypertrophied lenticels,46 which are enlarged lenticels that form on flooded roots up to immediately
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above the water table. These inflated lenticels aid in gas exchange and desiccate after flooding ceases.
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Collectively, the transport of VOCs in wood is not well quantified due to uncertainty in the phellem
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physiology. In studies involving CO2, the phellem’s resistance to diffusion, and therefore the flux, is
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variable, even for clones.47, 48
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MEASURING PHYTOVOLATILIZATION
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Direct phytovolatilization has been measured both in laboratory and field studies. In laboratory
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studies, plants are often dosed with contaminants, generally radiolabeled, in soil mesocosms or grown
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hydroponically. After a prescribed amount of time, plant parts are typically analyzed to perform a mass
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balance on the contaminant. The amount phytovolatilized may be directly measured by analysis of air
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passed through a sorbent trap in a closed system (preferable) or calculated from the amount of
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contaminant mass that remains unaccounted for in an open system. Note that the latter may be an
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overestimate due to incorporation of other loss mechanisms, such as phytodegradation. Additionally, if
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contaminant mass is captured on a sorbent trap, the phytovolatilization rate can be compared to
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elapsed time or to transpiration rate.
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At field sites, measurement of direct phytovolatilization from entire trees and other large plants
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is technically challenging and costly, so typically only individual leaves49 or, more commonly, branches
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are assessed. The leaves/branches are usually enclosed in a plastic bag, such as polyethylene,50 Tedlar51
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or Teflon-lined plastic52, or in a sealed solid plastic53 or glass chamber.54 The enclosure must not be
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permeable to the analyte of interest. To reduce humidity and to prevent the enclosed tissue from
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overheating, clean air is generally passed through the bag or chamber. The exiting gas is passed through 9
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a sorbent tube to capture the volatile analytes. Alternatively, a static system can be used that employs a
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cold trap or condenser to reduce humidity and can be sampled using a Summa canister.51 To measure
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the flux of VOCs emanating from tree trunks, a flux chamber is often used.55, 56 These chambers are
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sealed to the tree trunk and operate similarly to the bags that capture VOCs from leaves and branches.
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Clean air is passed into the chamber and contaminants are captured from the exit stream via a sorbent
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tube. An alternative technology for monitoring direct phytovolatilization is open path Fourier transform
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infrared spectroscopy (op-FTIR), a method that can detect VOCs in the air adjacent to the tree canopy.
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While FTIR has been attempted to monitor TCE volatilizing from a phytoremediation plot, the method
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was not sensitive enough to detect any TCE phytovolatilizing,49 although the method has been
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successfully used in laboratory experiments.57
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Phytovolatilization experiments with organic contaminants are not trivial to perform due to the
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volatility of the analytes of interest. Numerous problems with experimental setups have been
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encountered and discussed in the literature, both in laboratory and field setups.58, 59 See refs 35 and 60 for
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a discussion of challenges in monitoring biogenic VOC phytovolatilization. Perhaps most problematic is
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the need to seal the plant in a chamber while ensuring the plant undergoes natural photosynthesis and
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respiration. Artificial and natural light will heat the enclosed space while transpired water vapor will
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increase the humidity surrounding the leaves. The former can damage the leaf tissues while the latter
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can slow transpiration. To offset these effects, clean air is usually passed through the chamber at high
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flow rates to ensure humidity and temperature remain at acceptable levels. All the gas exiting the
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chamber must pass through a sorbent to collect volatile compounds. The sorbent must have high affinity
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for the targeted VOCs and low affinity for water. For many VOCs, Tenax meets these requirements. If
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VOCs are allowed to accumulate to high concentrations in the chamber, the concentration gradient
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across the plant boundary will lessen, likely resulting in lower diffusive fluxes and an underestimation of 10
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phytovolatilization. Additional difficulties often arise in sealing the experimental setup to prevent
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unmonitored release of VOCs. A particularly difficult region to seal is around the plant tissue. Stems are
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often sealed using closed cell foam, acrylic caulk or paraffin wax. In laboratory experiments, the
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tightness of system should be checked using wooden dowel rod or dead stem controls. However, a well-
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sealed experimental setup can also limit supply of necessary gases, such as oxygen to the roots, which
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may result in stress and increased root permeability. Care should be taken in plant selection, particularly
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when small tissue volumes are sampled. Stress or damage can cause stomatal closure and potentially
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affect both evapotranspiration and volatilization rates. Additionally, the dermal tissue of first-year wood
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(whips or branches) will not have the secondary dermal features present in older wood. Extrapolation of
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phytovolatilization rates from young woody plants to older woody plants is not recommended.
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Regardless of the system chosen, a suitable experimental setup must control temperature,
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relative humidity, and VOC concentrations inside the chamber, ensuring these values remain near
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ambient. If these values are not well controlled, substantial measurement error is likely. The specific
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flows of gases and other system parameters will depend on the plant, contaminant, and experimental
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system under study, so caution must be exercised when adapting systems from literature.
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REPORTS OF PHYTOVOLATILIZATION
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Laboratory Experiments
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Research regarding direct phytovolatilization has largely focused on the common groundwater
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contaminants, trichloroethylene (TCE) and tetrachloroethylene (PCE), using traditional
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phytoremediation plants such as willow (Salix sp.) and hybrid poplar (Populus sp. x P. sp.). These volatile
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contaminants are frequently reported to undergo phytovolatilization, although the degree of
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phytovolatilization varies substantially. In a closed system with hybrid poplars, Gordon et al.61 detected 11
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up by the plant was 70-90%. However, when a controlled field study was conducted, a minimal amount
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of TCE was phytovolatilized, as much of the TCE was dechlorinated in the subsurface. In another closed
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system experiment with poplar and willow cuttings dosed with 14C-PCE or 14C-TCE, 75% of the
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contaminant mass taken up by the plant was phytovolatilized (range: 27-96%, n=6), although
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experimental details are scarce.62 In flow-through reactors designed to provide a constant supply of
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contaminants, nutrients and oxygen, 90% removal of
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chloroform, although only ~1.5% of the removal resulted from phytovolatilization. The authors
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suspected much of the chloroform was degraded prior to phytovolatilization, but fate was not
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conclusively determined. Reid and Jaffé77 explored direct phytovolatilization of sulfur hexafluoride (SF6)
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in Typha latifolia and Scirpus acutus, finding daily fluctuations in the phytovolatilization rate. These
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fluctuations were correlated to temperature, photosynthetically active radiation (PAR) and vapor
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pressure deficit (VPD), which affected air-water partitioning, diffusivity and transpiration of this volatile
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compound. The correlation between VPD and phytovolatilization rate in Typha was thought to result
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from pressurized internal gas flows.32 Similar findings have been reported for methane direct
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phytovolatilization from Phragmites australis and Scirpus lacustris in constructed wetlands.78 Phragmites
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exhibited diel fluctuations in direct phytovolatilization, characterized by advective transpirative
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transport during the day and diffusive transport at night. Diel fluctuations in Scirpus were much less,
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implicating diffusive transport dominated, while transpiration-based fluxes were minimal. Direct
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phytovolatilization of methane from herbaceous plants in wetlands is relatively well studied (see
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reviews79, 80), as methane phytovolatilization can exceed methane volatilization from unplanted
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wetlands by an order of magnitude.79
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Direct phytovolatilization of methane from tree trunks has been measured less frequently than
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from herbaceous plants. Gauci et al.81 report methane fluxes of 6.1±1.2 µmol/m2/d in May, 97±88
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µmol/m2/d in June and 151±33 µmol/m2/d in October from wetland alder (Alnus glutinosa) trees (tree
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diameter 0.18-0.3 m; values are average ± standard deviation, n=3). The large fluxes of methane in May
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and June coincide with the tree in full leaf. Conversely, Terazawa et al.82 report relatively constant
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methane fluxes from Fraxinus mandshurica over five sampling events. Fluxes were measured at 263 17
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µmol/m2/d at 15 cm above ground surface, decreasing to 145 µmol/m2/d at 70 cm above ground
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surface, although this difference was not statistically significant (p=0.071). Pangala et al.83 measured
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methane fluxes of 25 – 277 µmol/m2/d in 7 of 8 tropical trees species. A multiple regression model could
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explain methane fluxes using stem diameter, wood specific density and porewater methane
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concentrations (R2 = 0.808, p
