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Background and literature review. Polycyclic aromatic hydrocarbons (PAHs) are a class of hydrocarbons that consist of at least two fused aromatic rings through two or more carbon atoms. PAHs are one of the major pollutants in urban areas, which can be dispersed in different environments: water, soil and sediments, air, plants and animals. Semivolatile organic compounds (SOCs) are distributed between the atmosphere and the surface of the earth. If consider that 80% of land surface of the earth is covered with vegetation, which exceeds the surface area of the land on which it is growing by 6-14 times, as well as count the fact that plant surfaces are covered with w°x or lipid layers, plants are important sinks for atmospheric SOCs, playing a significant rÐ¾le in their annual cycling. PAHs are airborne contaminants and present in the atmosphere in vapour and solid (particle-bound) phases. Due to their lipophilic properties, PAHs predominantly partition from the air to the leaf surface, as it mentioned above, which consist of nonpolar lipids and waxes.
The degree to which PAHs accumulate in the leaf depend on several factors:
Physico-chemical properties of PAHs. According to these properties, PAHs can deposit to leaves through a three basic deposition mechanisms: gaseous deposition, particle-bound deposition (wet and dry) and wÐµt dÐµposition of dissolved PAHs. Because of the hydrophobic nature of the PAHs the third deposition way is of low significance. High molecular weight non volatile PAHs with 5 or more rings tend to partition from the gas phase on atmospheric particles and then accumulate to plant surface, whereas low molecular weight volatile PAHs present in the atmosphere in a vapour phase and enter plant tissues primarily via open stomata by diffusion. Compounds of intermediate volatility with 4 rings are partitioned between gas and particulate phases depending on ambient temperature.
Partition of gaseous PAHs to leaf surfaces can be described as sorption process between vapour phase and leaf surface, where PAHs are transported from the ambient air to the vegetation until equilibrium is reached. The octanol-air partition coefficient (KOA) is an important parameter and helps to understand the partition of airborne pollutants to leaf surface. KOA is either measured directly or calculated from cÐ¾mpÐ¾und's air-water partition coefficient and octanol-water partition coefficient (KOW). Thus, accumulation of PAHs can be described with the log of KOA: (1) log KOA<~9 - equilibrium partitioning; (2) 9<log KOA<11 - kinetically limited gaseous deposition; (3) KOA>11 - particle bound deposition.
Kaupp and McLachlan (2000) found that 97% of particle-bound high molecular weight PAHs (with 5 and more rings) were attached to particles with aerodynamic diameters of <2.91 µm and only 1.2% of them were associated with particles with >8.6 µm aerodynamic diameter. They also found that with growing particle size PAH burdens constantly decreased.
Second factor which plays role in PAH deposition from the atmosphere is properties of the accumulating surface. There are great differences in the amount of adsorbed organic pollutants between various leaf surfaces which are exposed to the same air concentration. This can be explained by the different physiological features of leaves of different tree species, such as leaf morphology, lipid and epicuticular waxes content, stomata density and the presence or absence of hairs, which subsequently lead to a different efficacy in retaining airborne pollutants.
Total leaf area is the most essential plant feature, which affect the interception and accumulation rates of PAHs. Leaves contribute to the plant surface area much more than stems and twigs and c°n be used for comparison of accumulation rates between tree species. Jouraeva et al. (2002) suggested that the measuring the concentrations of PAHs per leaf area gives an accurate picture of the accumulation of PAHs rather than normalising them to per leaf dry weight, °s the leaves of diffÐµrent tree species may vary in the °mount of biomass per le°f °re°.
Howsam et al. (2000) proposed that the pubescent leaves tend to accumulate higher amounts of PAHs than hairless leaves. Thus, hairs were found to be effective at capturing of particles and preventing their washing-off. Barthlott and Neinhuis (1997) showed for the first time the interdependence between the surface roughness, caused by epicuticular wax crystalloids, water repellence and reduced particle adhesion. Hereby, the rougher the surface, the better the water repellence which is accompanied by reduced adherence of contaminant particles.
The cuticle is the outer continuous layer of the plant, which covers the aerial surfaces of the plants. The cuticle covers both adaxial and abaxial surfaces of leaves. It limits passive water loss and provides protection against some pathogens and minor mechanical damage. The rates of uptake of non-volatile lipophilic chemicals, which are deposited to leaf surfaces and cannot enter the plant via stomata by gaseous diffusion, are determined by cuticle. The plant cuticles are composed of the insoluble biopolymer cutin and waxes of various compositions. Wax which exists within the cuticle matrix is known as embedded wax or intracuticular wax, whereas wax that covers the cuticle matrix is referred to as epicuticular wax. The cutin was found to constitute 40-80% of the cuticle mass in some species.
There are great differences in the composition and morphology of epicuticular waxes between and within different plant species. Baker (1974) found that epicuticular waxes of brussels sprout (Brassica oleracea var. gemmifera) can develop different amorphous and crystalline forms, such as tubes, dendrites, filaments and plates. It is proposed that there is a strong connection between morphology and chemistry of epicuticular wax. Thus, the epicuticular waxes of most grass and Eucalyptus species can form two different morphology types, simple plate and tubular, according to the different dominating constituent of aliphatic components, primary alcohols (hexacosanol or octacosanol) and β-diketones (hentriacontan-14,16-dione or tritriacontan-12,14-dione) respectively. However, this is not always the case, e.g. plates can also be correlated with the presence of oleanolic acid in V. vinifera.
Epicuticular waxes are typically composed of n-alkanes, primary n-alcohols, n-aldehydes and fatty acids, where each class of compounds occur in homologous series with carbon atoms ranging from 20 to 40. In addition, it was identified that esters of fatty acids (C16-C34) and primary alcohols (C20-C36) can be formed, giving the compounds with the chain length of up to 70 carbon atoms. It was also established that n-alkanes, secondary alcohols, ketones, β-diketones and their hydroxy- and keto-derivatives show high preference for odd carbon numbers, whereas even carbon numbers predominate in primary alcohols, aldehydes, fatty acids and alkyl esters. The percentage of different individual compound classes may vary greatly. For example, leaf wax of H.vulgare was found to contain 89% of primary alcohols, 0.2-9.2% of alkanes, aldehydes, fatty acids and alkyl esters, whereas according to Bianchi et al. (1984) leaf wax of Zea Mays contained 9-42% each of above-listed class of compounds.
The amount of epicuticular wax may vary widely from 1µg/cm2 to several milligrams per square centimetre. It is possible to judge the thickness of the wax layer from its amounts assuming that the density of wax mixture is relatively constant and is about 0.8-1.0 g/cm3. In such case, 1µg/cm2 of wax would correspond to 10 nm of its layer thickness. It has been estimated for plants with 3-4 µm cuticle thickness the total amount of cuticular material may range from 180 to 1500 kg per hectar. This shows that plants play a considerable role in accumulating the lipophilic compounds in the environment. However, plants may only act as temporary sinks, as the leaves of deciduous trees are accumulated in the soil every year where it develops a lipophilic site for accumulation of pollutants.
Environmental conditions also influence the accumulation rates of PAHs on leaf surfaces. Ambient temperature influences the PAH-vegetation partition process. According to Simonich and Hites (1994) in spring and fall, when the ambient temperature is low gaseous PAHs partition into vegetation and at high ambient temperatures in summer some PAHs volatalize back to the atmosphere.
Leaf surface is surrounded by laminar boundary layer of air. Laminar boundary layer is characterised by almost absence of turbulence where the speed of wind is very insignificant. The thickness of laminar air depends on roughness of the leaf surface and speed of wind of upper air layers. It reduces at high winds and can be considerably thickened in calm weather. The atmospheric particles are trapped by laminar boundary layer and deposited on the leaf surfaces.
Barthlott and Neinhuis (1997) subjected water-repellent leaves to natural and artificial rains and found that particles of any size and different chemical nature were almost completely removed from the surface of the leaves until their surface waxy layers were not damaged and they explained it by high kinetic energy of the rain.
PAHs pose a risk to human health owing to their carcinogenic and mutagenic properties and therefore it is important to be able to monitor their concentrations in the environment and present both qualitative and quantitative analysis for these pollutants. Pollution of plants with PAHs is of a growing concern as they may enter a food chain starting with the vegetation and reach humans. It is also important fact that vegetation acts as an air filter and play role of a temporary reservoir for PAHs. Wagrowski and Hites (1997) estimated that 160 t of PAHs are deposited to plants every year and constitutes 4% of total emitted PAHs in northeastern region of the United States.
Another important point is that tree leaves are widely used as indicators to evaluate the atmospheric PAH levels in the urban areas. Such biomonitoring is found to be efficient and cost effective as it is easier to collect leaves than air samples.
PAHs are mainly produced by combustion which can be both natural (e.g. forest fires) and anthropogenic. Anthropogenic sources such as combustion in vehicles and domestic heating predominantly contribute to PAH emissions to the environment. PAH concentrations are higher in urban sites than in rural locations due to the closeness of pollution sources such as vehicles.
Aims and objectives
The aim of the present research project is to determine the extent to which leaves accumulate PAHs and investigate whether the leaves of different tree species accumulate these pollutants to different extents.
To achieve the aim of the project the following objectives were set:
To conduct leaf area and dry weight gravimetric measurements and determine the wax contents of the leaves.
To extract PAHs from leaf samples using ultrasonic extraction and conduct clean-up procedures of the extracts.
To identify individual PAHs both quantitatively and qualitatively by means of gas chromatography/mass spectrometry (GC/MS).
To compare total PAH concentrations per leaf areas and/or wax content of leaves of different tree species. Reveal any trends in PAH profiles and compare them with those in the literature.
The project aims to determine PAHs adsorbed on leaves (needles) of both deciduous tree species and evergreens. The concentrations of PAHs on leaves depending on seasonal variations or over a growing period will not be investigated due to the restricted time frame. Project will investigate the extents to which leaves of different tree species accumulate PAHs by comparing their PAH burdens (a mass per total leaf surface area and/or wax content).
16 USEPA priority PAHs, as well as 2- and 1-methylnaphtalenes, benzo[j]fluoranthene, benzo[e]pyrene and perylene were identified and quantified in leaves of 9 different tree species, which included:
Common holly (Ilex aquifolium)
Drooping juniper (Juniperus recurva)
Silver birch (Betula pendula)
English oak (Quercus robur)
Rhododendron (Rhododendron thomsonii)
Common Silver Fir (Abies alba)
Beech (Fagus sylvatica)
Sycamore (Acer pseudoplantanus)
Lime tree (Tilia europaea)
Study sites and sample collection
Two study sites within the city of Newcastle Upon Tyne were chosen. Back garden of the Drummond building of the Newcastle University and Moorbank botanical garden are situated next to the roads loaded with vehicular traffic (Fig. 3.1 and 3.2). Maps were modified from Ordnance Survey (2010).
Figure 3.2 Moorbank botanical garden
Leaves of different tree species were collected in September 2009 and May 2010 from both sites. Sampled areas are shown in Figures 3.1 and 3.2 as shaded circles. The exact locations of sampled trees were recorded using GPS navigator and aspects of the sampled sides of the trees were also identified (Table 3.1). All leaves were collected from 1.5-2 m height. Sampled tree leaves and needles were first air-dried for several days and then stored in paper envelopes until experiments commenced.
Extraction of PAHs and plant waxes
The solvent extraction was not exhaustive as epicuticular waxes (not embedded waxes) and PAHs on which they were adsorbed are of interest. The analytical procedure was tested several times prior to main analyses. There was a possibility that plant pigments such as chlorophylls and carotenes would be co-extracted along with epicuticular waxes and interfere with the analysis. There was also possibility that wax esters may cause peak overlapping on mass chromatograms. However, test procedures showed that plant pigments were retained by chromatographic column and there were no peak overlapping on mass chromatograms in the areas of interest. PAH levels on trial samples were roughly quantified in order to determine the amounts of internal and recovery standards that needed to be added.
Each leaf sample was weighed and appropriate amounts of 1,1'-binaphthyl solution in DCM was added as recovery standard directly to leaf samples prior to extraction using micro syringe (Table 3.3). After 30 minutes, when solvent was evaporated, leaves according to their sizes were placed into 250 ml or 500 ml beakers. DCM (200-400 ml) was then added to each sample in the beaker. Each beaker was placed into the ultrasonic bath for 1 minute. 3 consecutive extractions of each sample were made and extracts were then combined. After that, solvent volume of total extract was reduced to ~20 ml using rotary evaporator. The number of leaves which were subjected to extraction as well as their weights are given in Table 3.2.
Sample clean-up and gravimetric measurements
First, total extracts were subjected to filtering through a grade 1 (11 µm) filter paper to eliminate any present particles and clarify the extracts. After that, solvent was further evaporated from the filtered extracts using rotary evaporator to a volume of ~ 5 ml. Concentrated extracts were transferred to the 10 ml measuring cylinders and made up to 10 ml. A 1/9th aliquot of each sample was taken and transferred into the pre-weighed vials in order to determine the amounts of leaf waxes. The solvent was evaporated till dryness from aliquots using a stream of nitrogen gas. It was impossible to evaporate all the solvent only by using a stream of nitrogen gas as the solvent molecules were trapped within the leaf waxes. To remove all the solvent vials were placed in a warm bath (30-40°C). Vials were next weighed until constant weight and total amounts of extractable waxes were determined (Table 3.3).
The aliquots containing ~60 mg of extractable waxes or less were concentrated to a volume of ~0.25 ml and transferred into the vials with ~0.5 ml alumina. The remaining DCM was allowed to evaporate. The 500 mm long - 10 mm internal diameter glass columns were first plugged with cotton wool and then packed with silica and alumina (4:1). The aliquots which were adsorbed on alumina and free of DCM were loaded on top of the packed columns. PE (70 ml) was run through the columns to elute aliphatic hydrocarbons. PAHs containing fraction was eluted with DCM:PE (1:1) solvents mix (70 ml). Aliphatic fraction of each sample were concentrated down to a volume of ~3 ml and stored in capped vials. PAHs containing fractions were concentrated to a volume of ~1 ml and transferred to GC vials.
A procedural blank was also prepared by repeating all the analytical procedures but without leaf samples. A mix of standards was made by adding approximately equal amounts of 1,1'-binaphthyl, p-terphenyl and deuterated PAHs to a GC vial (Table 3.3).
A mix of deuterated PAHs and p-terphenyl solutions in DCM were added in appropriate amounts to each PAH containing fraction as internal standards prior to GC-MS analysis (Table 3.3). The mix of deuterated standards included 5 deuterated PAHs: naphthalene-d8, acenaphthene-d10, phenanthrene-d10, chrysene-d12 and perylene-d12.
The aromatic fractions of extracts, the standards mix and a blank sample were analysed by GC-MS, which was performed on Agilent 6890/7890A GC in split less mode, injector at (280°C) linked to a Agilent 5973/5975 MSD (electron voltage 70eV, source temperature 230°C, quadrupole temperature 150°C, multiplier voltage 1800V, interface temperature 310°C). The acquisition was controlled by a HP Compaq computer using Chemstation software in selected ion mode (30 ions 0.7 cps 35 ms dwell). The sample (1µl) in DCM was injected by an HP7673/7683B autosampler and the split opened after 1 minute. After the solvent peak had passed the GC temperature programme and data acquision commenced.
Separation was performed on an Agilent fused silica capillary column (30 m -0.25 mm internal diameter) coated with 0.25 µm dimethyl poly-siloxane (HP-5) phase. The column temperature was programmed from 50-310°C at 5°C/min and held at final temperature for 10 minutes with Helium as the carrier gas (flow rate of 1 ml/min, initial pressure of 50 kPa, split at 30 ml/min).
Peaks were identified by comparing the elution times of deuterated standards against corresponding PAHs as well as by recognizable peak patterns, elution order and relative retention times of PAHs.
Leaf area measurements
Surface areas of leaves were determined after extraction procedures by using AreaS 2.1 freeware (Samara State Agricultural Academy, 2005). Areas of any complexity can be calculated by comparing of two scanned images where the area of one of them is known. Thus, a scanned image of rectangular with known area of 5x5 cm2 was used to set up the scale. The areas of scanned leaves were estimated by the program automatically. Calculating error was no more than 0,001%. Calculated leaf area was doubled to obtain the total leaf surface area. Total surface areas (S) of needles were calculated by measuring the length (l), diameter (d) and the number (N) of needles extracted:
The total surface areas of leaves used in each sample are given in Table 3.4. The images of investigated leaves are given in Appendix 1.
Obtained GC-MS data was processed on Chemstation software (version E.01.01.335). Peak areas were integrated using RTE integrator and manual integration method. Unfortunately, some samples were contaminated with some polar compounds which were retained in the column and caused the tailing of the peaks in following samples. Contamination may come from either the detergents used in cleaning of the chemical dishes or contaminated solvents, alumina or silica as the experiments were carried out in a shared laboratory.
First, relative response factor (RRF) of internal standard (p-terphenyl) against surrogate standard (1,1'-binaphthyl) from the standards mix sample was calculated:
The chromatogram of standards mix can be seen from Figure 4.1 (Section 4). Recovery of 1,1'-binaphthyl was then calculated for each sample as following (Section 10, Table 10.4):
PAH amounts were determined by quantifying each PAH peak against the appropriate deuterated PAH (with the same number of rings) with the assumption that RRF of PAHs is equals to 1 (Table 3.5) as we do not know the exact amounts of analyte PAHs present in the sample:
, where x is the analyte PAH.
Estimated weights of the analyte PAHs were then corrected for recovery of surrogate standard (1,1'-binaphthyl) assuming that recovery standard reflects the conduct of the analyte PAHs:
Identification of PAHs
First of all, ion chromatograms of all added standards were obtained from standards mix sample. Figure 4.1 represents the combined ion chromatogram which was obtained by entering the commands in command line, which is on the main interface of Chemstation software. The algorithm of commands is given in Appendix 2. As it is known that deuterated PAHs come out just before their non-deuterated analogues, it was easily possible to determine the naphthalene, acenaphthene, phenanthrene, chrysene and perylene (Figures 4.2-4.5 and 4.8). Figures 4.6 to 4.9 illustrate combined ion chromatograms for all regions of interest, which were magnified between the particular time periods. Peaks were identified according to the elution order of the PAHs, recognizable peak patterns and relative retention times.
All samples were run in SIM mode and it was impossible to identify the peaks using program library. However, the library could not differentiate and identify the peaks within one m/z ratio, e.g. for ion 252 it gives equal percentage of matching (98%) for one peak with benzo[b]fluoranthene, benzo[k]fluoranthene, benzo[j]fluoranthene, benzo[e]pyrene and benzo[a]pyrene.
Table 4.1 summarises concentrations of individual PAHs and sum of 16 USEPA priority PAHs in 16 samples (D1-D7 and M1-M9). The PAHs which were quantified included 16 USEPA priority PAHs, as well as 2- and 1-methylnaphtalenes, benzo[j]fluoranthene, benzo[e]pyrene and perylene. The PAH concentrations are normalized to leaf surface area and expressed in ng per square decimetre. The raw data is given in Appendix 3.
Phenanthrene and anthracene, as well as benzo[b]fluoranthene and benzo[b]fluoranthene were quantified as a single peak due to their poor resolution.
The analysis of D5, D6 and D7 samples were performed to estimate the precision of the analysis. The mean concentrations of individual PAHs in these replicates are shown in Figure 4.10, with the standard errors shown as error bars.