Accepted Manuscript Title: Foliar Sorption of Emerging and Priority Contaminants under Controlled Conditions Author: Diana Calder´ on-Preciado V´ ıctor Matamoros Carme Biel Robert Save Josep M. Bayona PII: S0304-3894(13)00332-4 DOI: http://dx.doi.org/doi:10.1016/j.jhazmat.2013.05.016 Reference: HAZMAT 15103 To appear in: Journal of Hazardous Materials Received date: 15-2-2013 Revised date: 9-5-2013 Accepted date: 11-5-2013 Please cite this article as: D. Calder´ on-Preciado, V. Matamoros, C. Biel, R. Save, J.M. Bayona, Foliar Sorption of Emerging and Priority Contaminants under Controlled Conditions, Journal of Hazardous Materials (2013), http://dx.doi.org/10.1016/j.jhazmat.2013.05.016 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Transcript
Accepted Manuscript
Title: Foliar Sorption of Emerging and Priority Contaminantsunder Controlled Conditions
Author: Diana Calderon-Preciado Vıctor Matamoros CarmeBiel Robert Save Josep M. Bayona
Received date: 15-2-2013Revised date: 9-5-2013Accepted date: 11-5-2013
Please cite this article as: D. Calderon-Preciado, V. Matamoros, C. Biel, R.Save, J.M. Bayona, Foliar Sorption of Emerging and Priority Contaminantsunder Controlled Conditions, Journal of Hazardous Materials (2013),http://dx.doi.org/10.1016/j.jhazmat.2013.05.016
This is a PDF file of an unedited manuscript that has been accepted for publication.As a service to our customers we are providing this early version of the manuscript.The manuscript will undergo copyediting, typesetting, and review of the resulting proofbefore it is published in its final form. Please note that during the production processerrors may be discovered which could affect the content, and all legal disclaimers thatapply to the journal pertain.
20Highlights21Foliar sorption of organic micropollutants at two relative humidities is evaluated.22Rinsing following an incubation period was performed to evaluate the fate of contaminants on 23the leaf surface.24Neutral and basic contaminants were predominant in the leaf compartment.25Acidic compounds occur both in leaf tissue and rinsing water.26Relative humidity has a limited effect on leaf compartmentation.27
28
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Abstract29
Agricultural irrigation water contains a variety of contaminants that can be 30
introduced into the food chain through intake by irrigated crops. This paper describes an 31
experiment under controlled conditions designed to simulate sprinkle irrigation with 32
polluted water at two different relative humidities (40 and 90%). Specifically, shed 33
lettuce-heart leaves were spiked with an aqueous solution containing organic 34
microcontaminants, including pharmaceuticals (ibuprofen, diclofenac, clofibric acid,35
and carbamazepine), fragrances (tonalide), biocides (triclosan), insecticides (lindane), 36
herbicides (atrazine), phenolic estrogen (bisphenol A), and polycyclic aromatic 37
hydrocarbons (phenanthrene and pyrene). Following an incubation period (48 h), the 38
treated leaves were rinsed with water, and both the solution used to rinse them and the 39
leaves themselves were independently analyzed to investigate the foliar sorption and 40
uptake of the spiked organic contaminants through cuticle. The results showed that the 41
foliar sorption of emerging and priority microcontaminants in leaves wetted by 42
irrigation practices is related to their polarity (log Dow) and volatility (log kH), regardless 43
of their compound class and the relative humidity. The results thus underscore the need 44
to improve the quality of reclaimed water in crop irrigation, particularly when sprinkle 45
Water scarcity is a global challenge. Almost one fifth of the world’s population 56
lives in areas where water is physically scarce, and an additional 500 million people are 57
moving toward this situation [1]. It is widely accepted that climate change together with 58
the increasing variability of the agricultural sector each year due to economic conditions 59
will increase the difficulties and risks in the sector. This, in turn, could make 60
agricultural practices more vulnerable.61
Agriculture accounts for the bulk of the global water demand, consuming 70% 62
of all freshwater resources [2]. This demand will increase in the near future in 63
accordance with different climate-change scenarios and as a result of population growth. 64
Consequently, reclaimed water usage is emerging as a sustainable water resource in the 65
agricultural sector and one that might help to mitigate its vulnerability.66
However, reclaimed water, which is subject to state and local microbiological 67
and heavy-metal regulations, still contains organic microcontaminants [3], basically 68
because these contaminants are only partially removed or not removed at all by 69
conventional wastewater treatment plants (WWTPs). Hence, microcontaminants can 70
enter the hydrological cycle and reach the food chain through crop uptake [4, 5]. 71
Microcontaminants such as priority and emerging contaminants access plants via 72
three main pathways: (i) root and foliar uptake from aqueous solutions [6, 7]; (ii) vapor 73
uptake from the atmosphere [8]; and (iii) the deposition of contaminated soil and dust 74
on plant cuticles and subsequent diffusion of the contaminants through plant surfaces [9, 75
10]. 76
In green plants, gas and vapor exchange (O2, CO2 and H2O) mainly occurs in the 77
leaf surface. The stomata in the leaf provide a very effective way of regulating this 78
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exchange according to internal and external conditions [11]. Environmental stressors79
(primarily drought) and sunlight affect the stomata’s opening and closing mechanism, 80
and the plants’ survival depends on several factors, one of the most important of which 81
is its cuticle properties. The cuticle is the last barrier to prevent water loss and enable 82
the controlled exchange of abiotic and biotic material [12-14].83
Plant cuticles are solid-state lipid membranes made up of cutin, cutan, and 84
waxes that cover the aerial primary organs of higher plants. In addition to lipophilic 85
components, cuticles always contain small amounts of cellulose and pectins, but these 86
polar constituents are confined to the inner cuticular layer [15]. Nevertheless, the 87
coupling of stomata and cuticular defense mechanisms does not completely protect 88
leaves from the intrusion or loss of solutes, e.g. from nutrient leaching [16], foliar 89
uptake of non-volatile pollutants [17], crop protection agents [18], and fertilizers [19].90
Foliar uptake of microcontaminants is a complex process that depends on a 91
number of factors, such as the physicochemical properties of the compound and cuticle, 92
the surface area of the plant leaf, and environmental conditions. Among the most 93
significant factors affecting this uptake are the lipophilicity, molecular weight and 94
concentration of the microcontaminants on the leaf surface. Environmental factors such 95
as relative humidity can improve the cuticular penetration of hydrated ions by reducing 96
the hydrophobic properties of the cuticle surface, causing cuticle swelling, delaying 97
droplet drying, maintaining deposits in hydrated form, and/or redissolving salt deposits98
[20]. Chemicals in contact with foliage can partition onto the cuticle and be translocated99
to other plant compartments or diffuse into the plant through stomata, although the 100
relative importance of the latter pathway is still a matter of controversy [20, 21]. 101
Furthermore, chemicals can be taken up either through the hydrophilic pathway, which 102
consists of aqueous pores formed by carbohydrate strands submerged in the lipidic 103
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interfaces [15, 22], or the solid-state pathway [23]. Finally, it is worth mentioning the 104
transport capacity of the stomatal pathway due to its large size exclusion limit (10 nm –105
1m) under daylight conditions.106
In this study, an experimental set-up was designed and carried out to evaluate, 107
the final fate of selected microcontaminants including both emerging and priority 108
organic contaminants in the foliar compartment under simulated sprinkle irrigation. The 109
microcontaminants were selected according to their ubiquity in WWTP effluents and 110
physicochemical properties and ranged from highly hydrophobic to highly hydrophilic. 111
The experimental design consisted of two sets of shed leaves spiked with different 112
analytes at two relative humidities (RHs) to assess the importance of the 113
physicochemical properties of contaminants and environmental conditions in the foliar 114
uptake of contaminants. 115
116
2. Material and Methods117
118
2.1 Materials and Reagents119
Ibuprofen, carbamazepine, diclofenac, clofibric acid, tonalide, bisphenol A, 2,2’-120
dinitrophenyl, dihydrocarbamazepine, and triphenylamine (TPA) were purchased from 121
Sigma-Aldrich (Bornem, Belgium); trimethylsulfonium hydroxide (TMSH) and 122
triclosan were obtained from Fluka (Buchs, Switzerland); and fenoprop, lindane, 123
atrazine, phenanthrene, and pyrene were bought from Riedel-de Haën (Seelze, 124
Germany). Milli-Q water (R<18.2 MΩ cm-1, TOC <25 µg L-1, pH≈6.5) was used in all 125
the spiking experiments. 126
Florisil was ordered from Merck (Darmstadt, Germany). Anhydrous sodium 127
sulfate and sodium chloride were purchased from Fluka. Disodium hydrogen citrate 128
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sesquihydrate was bought from Aldrich (Milwaukee, WI). Trisodium citrate dihydrate 129
was bought from Sigma (St. Louis, MO). GC grade acetone, methanol, 130
dichloromethane, hexane, and ethyl acetate were purchased from Merck. The Na2SO4131
was baked for 5 hours at 450 ºC in a muffle furnace before it was used. 132
133
2.2 Experimental Set-up134
The experiment consisted in spiking shed lettuce leaves (ca. 90 cm2) with 135
aqueous solutions containing the target analytes and then incubating them at a set RH in 136
the darkness. First, the leaves were divided into two sets. The first was sprayed with 137
acidic compounds, while the second was sprayed with neutral and basic compounds 138
(Table 1). The sprayed solutions were prepared from stock solutions in Milli-Q water to 139
a final individual compound concentration of 4 μg mL-1, except for tonalide (0.24 μg 140
mL-1), phenanthrene (1.2 μg mL-1), and pyrene (0.14 μg mL-1) due to the lower 141
solubility. All leaves were identified, weighed, and had their petiole covered with 142
aluminum foil prior to the spraying. The spraying with the corresponding spiking 143
solution was performed on both upper and lower leaf surfaces until the drip point was 144
reached. Following the spiking, the excess solution was removed by gravity holding 145
leaves vertically, and each leaf’s weight was immediately recorded. Both leaf sets were 146
then split into two subsets: one was placed in a 40% RH incubation chamber, and the 147
other in a 90% RH chamber. The chamber dimensions were 40 dm3 (22 cm high x 35.5 148
cm wide x 51.5 cm long). The target RHs were achieved by placing a saturated solution 149
of MgCl2•6H2O in distilled water (for 40% RH) or distilled water (for 90% RH) [24] in 150
the blue tray (T) (Fig 1). Both leaf sets were incubated for 48 h in the corresponding 151
RH-controlled chamber. The exposure chambers were hermetically sealed to stabilize 152
the RH and avoid light exposure, thereby promoting stomatal closure due to darkness 153
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and making it easier to study cuticle permeability by decreasing the likelihood of 154
contaminant penetration via stomata and their photooxidation on the leaf surface. The 155
temperature and RH within the chambers were monitored frequently using HOBO H08-156
003 sensors (Onset, MA, USA) located in the chambers (Fig 1). The mean temperature 157
in the chambers throughout the experiment was 22.6 ± 0.5ºC, while the RH was either 158
40±2% or 94±12%, depending on the chamber. At the end of the incubation period, the 159
leaves exposed to contaminant solutions were retrieved, and five sets containing three 160
leaves each were formed within each subset (20 sets in total). Each set was immediately 161
rinsed with 200 mL of water and then carefully wrapped in aluminum foil. The water 162
used to rinse each group was recovered and stored in amber glass bottles. A third set of 163
leaves was used as a blank, following the above mentioned steps, using Milli-Q water as 164
the spiking solution and a 90% RH. The rinsed leaves were frozen until the sorbed 165
contaminant determination.166
167
2.3 Analytical Methodologies168
Rinse Water. The 200 mL recovered in the rinsing step were considered the 169
aqueous compartment. The determination of compounds in this compartment was 170
carried out following a previously described methodology [25]. Briefly, a sample 171
volume of 200 mL was spiked with 1.5 μg of a surrogate standard mix (i.e., fenoprop 172
for the acidic compounds, 2,2’-dinitrophenyl for polycyclic musks, and 173
dihydrocarbamazepine for neutral-basic compounds). Samples were percolated through 174
an activated polymeric solid-phase extraction cartridge (100 mg Strata X from 175
Phenomenex, Torrance, CA). The cartridges were eluted with 10 mL of the solvent 176
mixture hexane/ethyl acetate (1:1). The extract was evaporated to approximately 20 μL 177
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under a gentle nitrogen stream, and 246 ng of TPA were added as an internal standard. 178
The vial was then reconstituted to 300 μL with ethyl acetate. 179
Vegetable Tissue. Lettuce was extracted as previously reported [26]. Briefly, a 180
matrix solid-phase dispersion procedure was applied to the vegetable samples followed 181
by pressurized fluid extraction (Applied Separations, Allentown, PA) using 182
acetone/hexane (1:1) and two extraction cycles (13.5 min, 104 ºC, 110 bar). Finally, 183
TPA was added as an internal standard. 184
Target Analyte Determination. Methylation of the acidic carboxyl group (acidic 185
compounds) from both aqueous and vegetable samples was performed in the GC 186
injector port at 270 ºC by adding 10 μL of TMSH solution (0.25 mol L-1 in methanol) to 187
a 50 μL sample aliquot before injection. Obviously, this step was unnecessary for 188
samples from the neutral and basic sets. Both derivatized and non-derivatized samples 189
were injected onto a TRACE GC-MS (Thermo Scientific) in the electron impact mode 190
(70 eV ionization energy) fitted with a 20 m 0.18 mm ID 0.18 μm film thickness 191
Sapiens 5MS (Teknokroma, Barcelona, Spain). Helium was used as carrier gas 192
(99.9995% purity) at a constant flow rate of 0.6 mL min-1. The oven temperature was 193
held at 65 °C for 1 min, and then the temperature was programmed at 15 °C min-1 to 194
120 °C, at 6 °C min-1 to 160 °C, at 9 °C min-1 to 180 °C, at 6 °C min-1 to 220 °C, and 195
finally at 8 °C min-1 to 315 °C, holding the final temperature for 7 min. A volume of 1 196
μL of extract dissolved in ethyl acetate was injected in the splitless mode.197
Method Performance. GC-MS instrumental linearity for neutral and acidic 198
analytes ranged from 0.005 to 4.06 μg mL-1. The limit of detection (LOD) and limit of 199
quantitation (LOQ) for both water and vegetable tissue analysis were defined as the 200
mean background noise in a blank triplicate plus three and ten times, respectively, the 201
standard deviation of the background. For aqueous samples, the LODs and LOQs 202
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ranged from 0.002 to 0.087 μg L-1 and 0.003 to 0.075 μg L-1, respectively. For vegetable 203
tissues, the LODs and LOQs ranged from 6.6 to 33 μg kg-1 (fw) and from 7.6 to 40 μg 204
kg-1 (fw), respectively. Validation of method performance for vegetable tissue was205
achieved by means of evaluating the analyte recovery from a spiked matrix (0.1 μg g-1). 206
Recoveries and repeatability were calculated from triplicates. Recoveries ranged from 207
51% (carbamazepine) to 107% (diclofenac), while relative standard deviations ranged 208
from 9.5% (phenanthrene) to 23% (tonalide).209
210
2.4 Statistical Analysis211
All statistical analyses were performed using the STATGRAPHICS Centurion 212
XVI.I version 16.1.02 (VA, USA.) and SPSS 15 package (Chicago, IL). All data sets 213
were adjusted to have a normal distribution so as to enable parametric statistics. Normal 214
distributions were obtained using the Kolmogorov-Smirnov test. Mean comparisons 215
were followed by a one-way ANOVA and the Bonferroni correction method for 216
multiple comparisons, whereas correlation was followed by Pearson correlation 217
analysis. Multiple linear regression model analysis was performed to predict the 218
compound mass distribution between the lettuce leaves and rinse water (response 219
variable) according to the experimental data and physicochemical parameters220
(explanatory variables).221
222
3. Results and Discussion223
3.1 Contaminant Distribution between Leaf and Aqueous Compartments (LC-224
AqC) 225
Fig. 2 shows that neutral and basic (NB) compounds, namely, tonalide, lindane,226
atrazine, pyrene, and phenanthrene, and one weak acidic compound, namely, triclosan,227
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were predominant (>90%) in the leaf compartment (LC) at both RHs. The latter was 228
expected, since it has been documented that hydrophobic compounds are able to cross 229
leaf cuticles by dissolving and diffusing in lipophilic domains made up of cutin and 230
amorphous cuticular waxes [27]. Conversely, acidic compounds were partitioned 231
between the two compartments (leaf and rinse water). Though an apparently higher 232
mass of compounds was recovered from the aqueous compartment (AqC) at 90% RH233
than at 40% RH (Fig 2), no statistically significant differences (p<0.05) were found 234
between the treatments at different RHs for any of the studied compounds and by 235
compartment (LC and AqC). Although Schönherr [28] observed that high RHs enhance 236
the transport capacity of calcium salts by increasing the size and/or number of aqueous 237
pores, our findings suggest that stomata opening would have little bearing on the 238
transport of microcontaminants to the LC. A plausible explanation for this phenomenon 239
would be the unlikelihood of stomatal penetration by microcontaminants present in the240
aqueous solution due to the high surface tension of the sprayed Milli-Q water solution241
and the fact that the darkness of the experimental design promotes stomatal closure.242
243
3.2 Impact of Physicochemical Properties on Cuticle Sorption 244
The influence of some relevant physicochemical properties, namely solubility, 245
molecular weight (MW), molar volume (MV), volatility defined as the Henry’s law 246
constant (kH), and polarity (log Dow), on compound distribution (Dleaf/water) was assessed 247
by correlation and multiple linear regression modeling studies. In addition, water quality 248
could affect surface wettability and increase the leaf surface area exposed to 249
contaminated water. In this study, for simplicity’s sake, only Milli-Q water (pH=6.5) 250
was used, and therefore the acidic components according to the pKa values shown in 251
Table 1 occurred in ionized anionic form.252
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water
leafleafwaterleaf xVC
xFWCLogD
water
/
3.2.1 Correlation Studies253
First, the compound mass distribution between the leaf and rinse water (log 254
Dleaf/water) was calculated as follows (equation 1):255
256
(equation 1)257
258
where Cleaf and Cwater are the concentration of the specific analyte in the leaf and rinse259
water respectively. FWleaf is the fresh weight of the leaf and Vwater is the volume of the 260
rinse water (200 mL). The correlations were then stratified by RH, taking into account 261
the above-mentioned physicochemical properties (Table 1). Log Dow and Log kH262
displayed a positive correlation with log Dleaf/water at both RHs (p<0.05), whereas no 263
statistically significant correlations (p>0.05) were found for solubility, MW, or MV at 264
any RH (Fig. 3). These results suggest that the hydrophobicity and volatility of the 265
studied compounds are the most relevant physicochemical properties with regard to 266
explaining the accumulation of these compounds in the leaf. Furthermore, and as 267
mentioned above, no appreciable differences between RHs were observed. The positive 268
effect of hydrophobicity can be explained by either uptake or adsorption to the surface 269
of highly hydrophobic compounds [29], but the effect of loss due to volatilization at the 270
air-water interface cannot be ruled out. Such an effect would explain the positive 271
correlation with volatilization by reducing the concentration in the AqC. Nevertheless, it 272
may also favor transference to the stomata compartment and, thus, foliar uptake [30] not 273
relevant in the experimental set up used due to darkness.274
275
3.2.2 Multiple Linear Regression Model Studies 276
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Table 2 shows that log Dow and log kH were statistically significant with regard 277
to explaining the variability of the log Dleaf/water at 40% RH, whereas MW and MV were 278
also relevant at 90% RH. The positive value of parameter associated with MW may be 279
explained by adsorption on the leaf surface, whereas the negative parameter280
associated with MV may explain the leaf uptake of the compounds. It means that at 281
higher MW and lower MV adsorption and uptake are enhanced. It has been reported 282
that high RHs enhance the transport capacity of the penetration route of hydrophilic 283
solutes by increasing the size or number of polar pores [28]. Then, the size of a 284
molecule is not so critical to control the leaf entrance through these pores. The higher 285
accumulation of compounds with smaller MVs is consistent with a previous study [31],286
in which it was demonstrated that lipophilic and hydrophilic compounds with small 287
MVs, such as 2,4-D or bitertanol, are better able to diffuse through the leaf cuticle.288
The contribution of volatility in the model at 40% RH is higher than at 90%. 289
This is consistent with the higher water-air transfer at low RHs and the increase in 290
compound concentration in the drops deposited on the leaf surface due to higher water 291
evaporation rates. The effect of MW and MV at 90% RH may be explained by the 292
higher water coverage on the leaf surface throughout the study because water aids to the 293
compound distribution through the leaf surface and, consequently, to a better adsorption294
and leaf uptake.295
Altogether, log Dow has been demonstrated to play the most relevant role in the 296
assessment of the sorption of microcontaminants in lettuce leaves. As discussed above, 297
hydrophobicity may aid in the sorption of compounds into the leaf compartment. In 298
general, compounds of intermediate hydrophobicity (log Dow = 1-3) are taken up and 299
translocated more easily through the cuticle than compounds outside this range [32]. In 300
contrast, the large lipophilic surface of the plant foliage constitutes an ideal collector for 301
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hydrophobic compounds. Our results are consistent with the studies conducted by 302
Schreiber and Schönherr [29], who found that surface sorption and uptake of organic 303
chemicals into conifer needles increase with hydrophobicity.304
305
3.3 Irrigation Regimes and Food Safety Implications 306
Currently, all standards and guidelines for water reuse and agricultural irrigation 307
are primarily aimed at protecting health by controlling human exposure to pathogenic 308
organisms and a limited number of toxic chemicals [33]. The same is true of the 309
selection of irrigation technique, in which the most important factor taken into account 310
is meeting plants’ water requirements in an optimum manner, while minimizing the 311
sanitary risks intrinsic to any given irrigation technique. Indeed, little attention has been 312
given to the potential risks stemming from foliar sorption of organic microcontaminants 313
as a possible route to the human food chain and the ensuing repercussions for both the 314
environment and the population. Nevertheless, organic microcontaminants are found in 315
the low g L-1 range in reclaimed waters, and their uptake has already been documented 316
under real field irrigation conditions with surface and reclaimed waters [5, 7]317
In this regard, our findings suggest that foliar sorption of emerging and priority318
microcontaminants under sprinkling irrigation conditions is a relevant pathway for the 319
accumulation of hydrophobic microcontaminants in crop leaves. Furthermore, from 320
these results it can be inferred that washing lettuce leaves with water does not remove 321
microcontaminants. In conclusion, foliar sorption should be taken into account when 322
assessing actual human exposure and, thus, the risk posed by these microcontaminants. 323
The latter information is critical to making a more informed choice regarding the most 324
suitable irrigation regimes to minimize crop and, thus, human exposure to these 325
pollutants. 326
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4. Conclusions327
An experimental set-up aiming to evaluate the foliar sorption of selected organic 328
microcontaminants was carried out at two RHs. NB compounds were predominant in 329
the LC, while acidic compounds were partitioned between the leaf tissue and rinse330
water. No statistically significant (<0.05) differences were found between RHs with 331
regard to mass distribution between the two compartments for any compound. However, 332
the impact of each compound’s physicochemical properties on its final compartmental 333
distribution was shown through the use of correlation and multiple linear regression 334
model analyses. These statistical analyses showed that volatility and polarity play an 335
important role in the final fate of the target compound and its plant uptake potential.336
This study has demonstrated that foliar sorption of emerging and priority 337
microcontaminants under simulated sprinkle irrigation conditions is feasible and offers 338
insight into their sorption in crop leaf cuticles. Although, care should be taken in 339
extrapolating the results obtained from shed leaves, our conclusions could be useful for 340
garnering further insight into the foliar uptake of organic microcontaminants by crops in 341
fields irrigated with reclaimed water and for minimizing population and environmental342
exposure.343
344
Acknowledgments345
This study was funded by the Catalan Food Safety Agency (ACSA) and the 346
Spanish Ministry of Economy and Competitiveness (CGL2011-24844). Dr. V. M. 347
would like to acknowledge a JAE-Doc contract from the CSIC, and to thank the 348
European Social Fund and the Consorci per a la Defensa de la Conca del Besòs (CDCB) 349
for the WWTP use.350
351
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Table 1. Relevant physicochemical properties of the selected compounds.440Compound Physicochemical properties
Chemical structure
Solubility (mg L-1)
Molecular weight (Da)
pKaMolar
volume, (m3 mol-1)
Henry Law constant
(dimensionless)Log Dow*
Acidic compoundsIbuprofen
41.1 206 4.4 182.1 6.2 x 10-6 1.10
Diclofenac
17.8 2954.0
206.8 1.94 x 10-10 1.17
Triclosan
17.0 289 7.9 194.3 2.04 x 10-7 4.61
Mecoprop
620 2143.1-3.8 169.6 7.45 x 10-7 -0.86
Clofibric acid
583 214 3.46 169.5 8.96 x 10-7 -1.32
Neutral and basic compoundsCarbamazepine
17.7 236 13.4 186.6 4.42 x 10-9 2.25
Bisphenol A
300 228 9.5 199.5 3.74 x 10-10 3.64
Tonalide
0.24 258 N 280.8 1.73 x 10-3 5.80
Lindane
10.0 290N
182.6 1.05 x 10-2 4.26
Atrazine
30.0 215 N 169.8 1.83 x 10-7 2.82
Pyrene0.14 202
N161.9 3.39 x 10-4 4.93
Phenanthrene
1.29 178N
157.6 2.10 x 10-3 4.35
N: neutral. *Dow was calculated as follows 441
442
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443Table 2. Multiple linear regression model to describe the relationship between log Dleaf/water and 5 444physico-chemical properties.445
446Log Dleaf/water
40% RH 90% RHPhysico-chemical property
β value p-value β value p-valueSolubility 0.0016 0.0565 0.0015 0.1626
Figure 1. Experimental set-up depicting the incubation conditions and different treatments of 546the heart’s leaves (NB, neutral & basic compounds). See Table 1 for compound identity and547physico-chemical properties. Incubation chamber dimensions were 40 dm3 (22 cm high x 35.5 548cm wide x 51.5 cm long). The target RHs were achieved by a saturated solution of 549MgCl2•6H2O in distilled water for 40% RH and distilled water for 90% RH (Young, 1967),550contained a blue tray (T). Real-time Temperature and RH were in situ measured and stored in a 551data logger both located underneath (D).552
Figure 2. Final analyte distribution between the aqueous and leaf compartments 604following the incubation period of 48 h: a) 40% RH and b) 90% RH in the incubation 605chambers shown in Fig 1.606
Figure 3. Compound mass distribution between leaf and rising water (log Dleaf/water) 643plotted against the polarity (log Dow) and volatility (kH) at 40 and 90% RH respectively.644
