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Bioconcentration Factors for Volatile Organic Compounds in Vegetation

Michael H. Hiatt
U.S. Environmental Protection Agency, National Exposure Research Laboratory
Environmental Sciences Division. P.O. Box 93478, Las Vegas, Nevada 89193-3478

  Phone: 702 798 2381. Fax: 702 798 2142. 
E-mail: hiatt.mike@epa.gov.
[Note:  minor content and formatting differences exist between this web 
version and the published version]

Table of Contents

Abstract

Samples of air and leaves were taken at the University of Nevada-Las Vegas campus and analyzed for volatile organic compounds using vacuum distillation coupled with gas chromatography/mass spectrometry (VD/GC/MS). The data were used to estimate the bioconcentration of volatile organic compounds (VOCs) and to characterize the equilibration of VOCs between the leaves and air.

The bioconcentration of volatiles in the leaves of some species can be predicted using the partition coefficients between air and octanol (Koa) and only considering VOC absorption in the lipid fraction of leaves. For these leaves, the bioconcentration factors agreed with existing models. Leaves of some species displayed a bioconcentration of volatiles that greatly exceeded theory. These hyper- bioconcentration leaves also contain appreciable concentrations of monoterpenes, suggesting that a terpenoid compartment should be considered for the bioconcentrationof organic compounds in leaves. Adding an additional "terpenoid" compartment should improve the characterization of volatile organic compounds in the environment.

The uptake of VOCs from air by leaves is rapid and the equilibration rates are seen to be quicker for compounds that have higher vapor pressures. The release of VOCs from the leaves of plants is slower for hyper-bioconcentration leaves.

Introduction

Volatile organic compounds (VOCs) are an important group of chemicals that permeate our environment. The concentrations of VOCs in vegetation is one factor that must be considered for an assessment of the enviroment's exposure to these chemicals. The major route for plant uptake of volatile hydrophobic compounds is through sorption of the compounds directly from air.1 This uptake by vegetation is suggested to be species-dependent.2 Several models are used to predict bioconcentration factors based on the partitioning of organic compounds between air and an organic phase (as octanol) and between air and an aqueous phase.3,4,5 Documented determinations of VOCs in vegetation resulting in bioconcentration factors (BCF) are limited to tetrachloroethene in pine needles4 and 1,2,4-trichlorobenzene in soybeans.6

The uptake of tetrachloroethene in pine needles is reported to be more complex than the published model.4 In that study, pine needles in a chamber were exposed to elevated levels of tetrachloroethene (PCE), and the concentration of PCE in the needles was predicted by its Koa and lipid content. When the needles had been exposed to the much lower envirornmental levels, the concentration of PCE was much greater than predicted. It was suggested that an additional compartment in the needles bioconcentrated PCE in excess of theory, but had a limited capacity to absorb PCE.

This report is an investigation of the behavior of volatile organic compounds (VOCs) and tests the use of lipid content and Koa to predict BCF in an uncontrolled environmental setting. The urban environmental variations encountered in these experiments introduce errors in estimating BCFs but highlights a major factor affecting the equilibria of VOCs between leaves and air.

The environmental site investigated for this study was at the U.S. Environmental Protection Agency's National Exposure Research Laboratory on the University of Nevada-Las Vegas (UNLV) campus. The VOCs monitored in this study were those that are persistent in the environment, tended to bioconcentrate in an organic phase (hydrophobic), and could easily be detected in air and leaves. Nine different plant species found on the UNLV campus were analyzed using vacuum distillation coupled with gas chromatography/mass spectrometry (VD/GC/MS) to determine concentrations of VOCs. Grab air samples were taken before and immediately after collection of leaves to determine the concentration of VOCs in air.

Experimental Section

Vacuum Distiller. The appartus modified with a cryogenically cooled condenser (Associated Design & Manufacturing Company, Alexandria, Virginia), has been previously described.5 A pirani vacuum (Edwards Model 1000) gauge was placed at the vacuum pump to monitor the integrity of the apparatus under vacuum. The temperature of the sample chamber six-port valve was maintained at 150 C (Valcon E rotor). All transfer lines were heated to 90 C. The Nupro toggle valves were replaced with Peter Paul Series 70 Model 72 solenoid-action valves (Peter Paul Electronics Co., Inc., New Britain, CT).

The vacuum distiller was modified for a helium sweep of the condenser column to remove condensate between vacuum distillations. A helium transfer line (5 psi) was connected at the top of the condenser column (Nupro toggle valve), and a helium exit vent (Nupro toggle valve) was attached to the transfer line between the sample chamber valve and the condenser.

The condenser column was normally held at 7.5 2.5 C during vacuum distillations and at 90 5 C between distillations.

GC/MS Apparatus. A Hewlett-Packard mass spectrometer (Model 5972) and gas chromatograph (HP5890 Series II with Model MJSC metal jet separator) with a 60-m x 0.53-mm i.d., 3.0-m film thickness, VOCOL capillary column (Supelco, Bellefonte, PA) was used for the determination of analytes from the vacuum distillation apparatus. Gas chromatograph operating conditions were: 3 min at 10 C; 50 C/min ramp to 40 C; 5 C/min ramp to 120 C; 20 C/min ramp to 220 C; and isothermal at 220 C for 3.4 min, resulting in a total run time of 28 min. The jet separator was held at 210 C and the transfer line at 280 C. The mass spectrometer was set to scan from 38 to 270 amu at 3.1-s. The injector was interfaced to the vacuum distillation apparatus by connecting the carrier inlet gas line to the cryoloop valve and then back to the injector. The injection or inlet temperature was 240 C and the inlet pressure was 10 psi.

The mass spectrometer was also operated in the selected ion mode (SIM with 100 msec dwell on each ion being monitored. The additional sensitivity gained by using SIM became necessary to consistently detect and measure the analytes. The ions monitored for the analytes, surrogates, and monoterpenes are listed in Table 1.

Leaf Analysis. Samples were prepared using 10 g of fresh leaves removed from their stems and placed in the sample vessel. There were no attempts to mince or mix the tissue. The samples were spiked with 10 L methanol containing surrogate compounds directly in sample vessels, (a 100-mL round bottom flask fitted with a 15-mm O-ring connector) which were also used to contain the samples during both vacuum spike and vacuum distillation.

The vacuum spike was performed as previously described for 10-g fish samples.7 The vacuum spike equilibrates a spike with the sample matrix in a vacuum at ambient temperatures within 3 hours. For this project, the vacuum spikes were performed overnight, and the samples were analyzed by VD/GC/MS the following day.

Before analysis, the vacuum distiller's condenser column was cooled to 7.5 C to condense water evaporated from the sample during distillation. The sample chamber containing the spiked sample (room temperature) was distilled under vacuum for 5 min. Water vapor was collected on the condenser column while the fraction containing the analytes was collected in the cryoloop immersed in liquid nitrogen (-196 C). The sample chamber valve was closed at the completion of the vacuum distillation and the cryoloop valve switched to allow the GC carrier gas to sweep the cryoloop and pass to the capillary column. The cryoloop's liquid nitrogen bath was removed, and the cryoloop temperature was ballistically heated to 120 C to volatilize the distillate. After transfer of the distillate to the GC was complete (3 min), the cryoloop valve was again switched and the cryoloop was heated to 200 C and then allowed to cool to room temperature. After the sample was vacuum distilled, the condenser column was heated to 60 C, and the condenser flushed with nitrogen gas, while the nitrogen gas/condenser valve and vent valve were opened to remove most of the trapped material. After 3 min the condenser-nitrogen gas line and vent valves were closed, and the system was evacuated with the vacuum pump for an additional 10-min period to remove any condensed water and contaminants remaining after the nitrogen gas flush.

The concentrations of analytes in leaves were determined using the surrogate-based matrix correction technique.7,8,9 For this study, the recoveries of analytes from the matrix were compared with the surrogates respective to their relative volatility between octanol and air (aKoa). With the analyte recovery prediction, their responses were corrected for matrix effects and then the concentration was determined.

Three groups of surrogates were used. The first group of surrogates was used to determine the matrix effects for analytes with aKoa values below 5000. The surrogates in the first group were fluorobenzene, 1,4-fluorobenzene, toluene-d8, and 1,2-dibromoethane-d4. The second group of surrogates was used to predict recoveries for those analytes with aKoa  values between 5000 and 22000. The second group's surrogates were 1,2-dibromoethane-d4, bromobenzene-d5, and 1,2-dichlorobenzene-d4. The third group of surrogates was used to predict the recovery of analytes with Koa greater than 22,000. This group of surrogates was 1,4-bromofluorobenzene, 1,2-dichlorobenzene-d4, 1,2,4-trichlorobenzene-d3, and naphthalene-d8. The list of aKoa values is presented in Table 1.

The concentrations of volatile organic compounds were determined relative to the dry-weight of the leaves. The dryweight for leaves was determined as the weight after heating at 115 C overnight. The samples were dried in sample vessel after their vacuum distillation. The sample dry-weight was used to determine the BCF.

Leaf samples of grass (mixed), ivy (Hedera canariensis), olive (Olea eurpaea), strawberry tree (Arbutus unedo), Chinese photina (Photina serrulata), mock orange (Pittosporum tobira), holly (Ilex cornuta burfordii), pine (Pinus eldavica), and juniper (Juniperus sabina tamariscifolia) were analyzed.

Air Analysis. Air samples were collected using a 1- L Erlenmeyer flask modified with an O-ring connector (15 mm). The Erlenmeyer flask was sealed using a cap made from a stainless steel O-ring connector (15 mm) attached to a stainless steel toggle valve with Swagelock fittings (Nupro 4BKT). Before sample collection the flask and valve/cap were attached (1/4 in. Swagelock fittings) to a port on the vacuum distillation apparatus and evacuated. An air grab sample was then collected in the flask (disconnecting the O-ring connection), adding the 5-L surrogate solution and then reconnecting the valve/O-ring cap. The flask assembly was then reattached to the vacuum distiller for analyses. The air sample was evacuated from the flask (open cap-toggle valve), passed through the vacuum distiller condenser and focused in the system cryotrap (-196 C). The condenser was held at 90 C during the sample transfer. After five minutes of collecting the analytes in the cryotrap, the valving was switched to desorb mode, and the cryotrap heated to 100 C, transferring the focused material to the GC/MS. The determination of analytes in the air samples was performed using external standard quantitation. Results were not corrected to STP.

Results and Discussion

The VOC analytes for this study were selected based on their frequency of occurrence in air, ability to bioconcentrate, and freedom from mass spectral interferences. The analytes that met these criteria were monitored in the air and vegetation as a means to understand the bioconcentration of VOCs and their potential use as a measurement of environmental quality. Polar analytes were not investigated as they were not expected to appreciably bioconcentrate.

The vacuum distillation of vegetation also extracted compounds that introduced mass spectral interferences for some compounds thatwere easily detected in air. Chloroform and 1,1,1-trichloroethane were too low in concentration to be distinguished from coeluting compounds that were present in all leaf samples, and therefore the BCFs for these analytes were not determined. The compound, o-xylene, co-eluted with the monoterpene, tricyclene. This co-elution with tricyclene resulted in interferences for all the o-xylene mass fragments and negated measurement of o-xylene in leaves that contained monoterpenes. The o-xylene isomers, m-xylene, p-xylene, and ethylbenzene were easily detected.

The relative volatility factors for water and organic phases were experimentally determined for monoterpenes using previously described techniques.7,9 The relative volatilities between water and air for the monoterpenes were experimentally determined to be between 2 and 5. The relative volatilities between a octanol and air for the monoterpenes were experimentally determined and presented in Table 1. The monoterpenes display a great affinity for an organic phase and behave very similarly to the analytes in this study. The monoterpene compounds listed in Table 1 were used to quantify the fraction of monoterpenes nearest to it and using the surrogate corrections as previously described. Total monoterpene concentrations reported in Table 2. These concentrations are estimates because the large amount of vacuum distilled monoterpenes exceeded the calibration (for pine, juniper, and rosemary) and were not chromatographically resolved.

The concentration of the organic compounds in the air fluctuated during the day. Generally the concentration of compounds in the air was highest in the morning, dropped rapidly during the day to the lowest concentration at midday, and rose in late afternoon. These concentrations were stable in the morning until the air began to mix (typically between 7:00 a.m. and 9:00 a.m.). The higher concentrations of analytes in the morning reflect the denser (colder) morning air temperatures, and a lack of thermal mixing currents. An increase in aromatic hydrocarbons was observed to coincide with early morning and late afternoon automotive traffic flows. The concentrations of analytes were generally stable between 11:00 a.m. and 3:00 p.m. With warmer temperatures at midday and the activity of thermal air currents, the lower morning air is mixed to higher elevations and becomes more dilute.

Using toluene as an example, Fig. 1 shows how the morning and afternoon air concentration of compounds varied during this study. As the concentration low at midday reflects mixed air, the consistency of the midday concentrations suggests the total amount of these compounds in the Las Vegas Valley was constant during the period. It was also observed that the morning analyte concentrations tend to decrease from winter to summer.

The variation of VOC concentrations in leaves during a day was investigated by comparing daily morning concentrations (7:30 to 8:30 a.m.) and afternoon concentrations (1:00 to 2:30 p.m.). The percentage drop in the concentration of analytes in leaves was compared with the percentage drop in the concentration of analytes in the air. These data are presented in Fig. 2 with the percent change in leaves divided by the percentage change in air being reported as relative change and compared to the vapor pressure of each analyte. A relative change equal to 100% suggests that the concentration of the analyte changes in a plant leaf and in the surrounding air at the same rate and that equilibration (by release of VOCs by leaves) between the leaf and air is rapid (less than 5 hours). It is also shown that the vapor pressure of a compound is closely related to how quickly the compound equilibrates between the leaf and air. This effect was also seen when spikes were being equilibrated with fish tissue in a vacuum.7

The rate of equilibration of an analyte between a plant's leaves and air (Fig. 2) is strongly dependent on species. This figure shows that while the afternoon may be a good sampling period for air (stable concentrations), the content of VOCs in leaves, however, are unlikely to reflect equilibration with the air. The lower an analyte's vapor pressure during sampling the less likely it is to reflect equilibration during afternoon sampling. Fig 2 also indicates that as monoterpene content in leaves increases, the release of VOCs slows hindering equilibration. Sampling in the afternoon, therefore, was not considered appropriate for the determination fo BCF.

It has been reported that the bioconcentration between air and leaves is affected by plant characteristics by impacting chemical transfer times between the air and leaf.3 The rate leaves take up a chemical was reported to be greater than the release rate and the ratio of the rates equivalent to the leaf's BCF for the chemical.10 Therefore it is more likely that the VOC content in leaves reflects equilibrium when air concentrations are increasing, rather than decreasing. Equilibration becomes more favorable if air concentrations are stable following an increase in temperature. These criteria were best met by early morning conditions. Only leaves that were collected in the morning were used to determine the BCF values reported in this study.

The BCFs for VOCs in the different leaves were calculated using the formula

 BCF = C1/Ca     (1)

where C1 is the concentration of the compound in leaves, and Ca is the concentration of the compound in the air and concentrations are in mass/volume units. The concentrations in the plant leaves were determined on a dry-weight basis and made equivalent to volume by assuming the density as 1 (1g leaves dry-weight = 1 mL).

For analytes that are lipophilic, BCFs are closely related to Koa and can be considered as the partitioning of analytes between air and an organic phase, conventionally octanol. The partitioning of a compound between liquid (octanol) and gas phases is described as

K1g = dLRT/grM    (2)

where dL is the density of the liquid phase, R is the universal gas constant, T is temperature in Kelvin, and g and r are the activity coefficient and the vapor pressure of the compound, respectively, and M is the molecular mass of the liquid phase.11

Changes in system temperature have a significant impact on the vapor pressures of VOCs and an insignificant impact on activity coefficient for lipophillic compounds dissolved in a lipophillic phase. BCF would be affected by changes in temperature as Klg. Comparing T/r at 20 C to T/r at the sampling temperature generates a correction factor for reporting BCF for 20 C. The experimental values for BCF were all standardized for 20 C and reported as BCF20. Vapor pressures of the analytes at sampling were estimated.12 The BCF20 values by analytes and type of plant are presented in Table 2.

One problem encountered was the air present in the headspace of the sample vessel containing samples. For instance, air in the 100-mL volume of the sample vessels would contain the same amount of an analyte as 2 g of grass (dry weight) if the bioconcentration factor was 50. While the headspace was not a critical problem for most analytes in leaves (BCF > 200), it could be a source of bias. If precautions were not made, a sample would have a headspace contribution at two separate times. The first contribution would occur during sample collection as the sample headspace would contain analytes present in the air during collection. A second contribution occurs when the sample is removed from the vacuum spike apparatus and the vacuum in the sample vessel is refilled with laboratory air. The contribution from the "second headspace" was minimized by filling the vacuum spike apparatus and sample vessel with nitrogen before a quick transfer of the sample to the vacuum distillation apparatus. The concentrations of analytes in plants were corrected for the contribution of analytes from the headspace.

The determination of BCF could be biased by morning sampling if equilibration of VOCs between the leaves and air was not reached overnight. A high bias would occur when the concentrations of VOCs in the morning air were lower than the previous morning and the release of VOCs by leaves during the 24 hour period was too slow to reach equilibration. A low bias would occur when VOC concentrations in the morning air increased faster than uptake of VOCs by leaves. Because the uptake is quick, it was not considered that the concentration of the leaves at morning would normally be less than that reflecting equilibration. However, an atypical rapid increased of VOCs from automobile exhaust just prior to air sampling would produce a low bias. During the study no attempts were made to identify or eliminate outliers as a potential result from one of these biases. Such biases would be expected to contribute to the variations in the determination of BCF values. Variations in BCF20 that reflect natural or seasonal changes in leaves were expected to be important but could not be diStinguished during the period of this study.

The BCF20 values for the compounds in the leaves of the different plant species are averages of the multiple BCF20 determinations and are reported in Table 2. These BCF20 values were then compared with that predicted by accumulation of compounds in a lipid phase as Koa. Figure 3 presents the concentration of naphthalene in mock orange (dry-weight) and in air (morning), and the calculated BCF20 for the period of this study (between January 2, 1997 and June 23, 1997). The daily BCF20 was shown to vary during the period and assumed to reflect errors associated with using the dynamic environmental conditions.

Leaves from grass, ivy, holly, oleander, olive strawberry tree, and Chinese photina were found to have low levels of monoterpenes and they also contained similar concentrations of the monitored analytes. These leaves were found to take up chemicals from the air and could be described using the following formula

BCF = 0.19 + 0.7Kwa + 0.05Koa    (3)

where 0. 19 is the fraction of the plant (wet weight) that is air, 0.7 Kwa is the water-air partition coefficient multiplied by the plant's water content, and 0.05 Koa is the octanol-air partition coefficient multiplied by the plant's lipid content.3 For this study, the air and water phase terms are not significant and therefore the BCF is determined based on the lipid fraction. The fraction of the plant considered lipid, however, can vary. Some of the BCF determined were larger than a lipid content of 5% wet weight would provide and therefore lipid contents could be greater than 5% but with the limitation that lipid content could not exceed the dry weight of a plant. There were no attempts to measure the lipid content of the leaves. The leaves that could be considered as conforming to equation 3 and having a lipid content up to the plants dry weight would follow the following equation

BCF20 = C1/Ca = Xo/Koa   (4)

where Xo is the fraction of the plant that behaves as lipid and has a value between 5% wet weight and the % dry weight.

Of the nine plant species studied, the leaves of mock orange, pine, rosemary, and juniper were found not to conform to eq 4. These leaves had VOC concentrations that required a lipid fraction that exceeded the plant's dry weight. These plants also had significant concentrations of indicating a terpenoid phase (including properties of plants that manufacture terpenes) was responsible. The VOC partitioning between this terpenoid phase and air would have to be greater than Koa. An expansion of eq 3 is recommended as follows:

BCF = Va + VwKwa + V1Koa + V1Kta     (5)

where Va, Vw, Vl, and Vt, are the volumes of plant leaf that are respectively, air, water, lipid, and terpenoid. Kta is the partition coefficient between the terpenoid phase and air. A limited capacity of this volume for uptake of a chemical would be consistent with the reported compartment (and Kta greater than Koa for tetrachloroethene) in pine needles that had to be saturated before eq 4 was valid.4 Measurement of the volume Vt and the Kta (or activity coefficients for terpenoid phase) was not performed.

A simplification of eq 5 provides the following:

BCF = 0.19 + 0.7Kwa + (0.05 + P)Koa   (6)

where P is equal to the product of Vt times the activity coefficient of a chemical in octanol divided by the activity coefficient of the chemical in the terpenoid phase. The experimental determinations of BCF can be used in eq 6 to solve for P. The resulting values for P are presented in Table 3. It should be noted that P is only valid when the chemical concentration in air is such that the terpenoid capacity is not exceeded. It is likely that values for P will vary with seasonal changes in leaves that alter the terpenoid phase.

Conclusion

The uptake of VOCs by vegetation can be much greater than currently estimated. An additional uptake due to a terpenoid phase can exceed predictions by an order of magnitude. If not considered, this factor could introduce significant errors in modeling ecosystems and the fate of VOCs. Additional investigations as to the nature of the terpenoid phase and its impact on the uptake of other chemicals should be conducted.

Acknowledgment

The EPA, through its Office of Research and Development (ORD), funded and performed the research described here. It has been subjected to the Agency's peer review and has been approved as an EPA publication. Mention of trade names or commercial products does not constitute endorsement or recommendation for use.

References

  • Travis, C. C.; H. A. Hattemer-Frey, Chemosphere 1988, 17, 277-283.
  • Buckley, E.H., Science 1982, 216, 520.
  • Paterson, S.; Mackay, D.; Bacci, E.; Calamari, D., Environ. Sci. Technol. 1991, 25, 866-871.
  • Frank, H.; Frank W., Environ. Sci. Technol. 1989, 23, 365-367.
  • Paterson, S.; Mackay, D.; McFarlane, Environ. Sci. Technol. 1994, 28, 2259-2266.
  • Paterson, S.; Mackay, D.; Gladman, A., Chemosphere 1991, 23, 539-565.
  • Hiatt, M., Anal. Chem. 1997, 69, 1127-1134.
  • Hiatt, M., Farr, C., Anal. Chem. 1995, 67, 426-433.
  • Hiatt, M., Anal Chem. 1995, 67, 4044-4052.
  • Bacci, E.; Cerejeira, M.; Gaggi, C., Chemello, G.; Calamari, D.; Vighi, M., Chemosphere 1990, 21, 525-535.
  • Ioffe, B.; Vitenberg, A. Head-Space Analysis and Related Methods in Gas Chromatography; John Wiley and Sons: New York, 1984; p. 14.
  • Lyman, W.; Reehl, W.; Rosenblatt, D. Handbook of Chemical Property Estimation Methods; American Chemistry: Washington DC, 1990; Chapter 14, 12-15.

 

 

 


 

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