Phthalate Esters (Handbook of Environmental Chemistry)

The Handbook of Environmental Chemistry Vol. 3, Part Q (2003): 1– 7 DOI 10.1007/b11460

Introduction Marian K. Stanley 1 · Kenneth A. Robillard 2 · Charles A. Staples 3 1 2 3

American Chemistry Council, 1300 Wilson Blvd., Arlington, VA 22209, USA E-mail: [email protected] Eastman Chemical, Rochester, NY, USA Assessment Technologies, Inc. 10201 Lee Highway, Suite 580, Fairfax, VA 22030, USA

The esters of 1,2-benzene dicarboxylic acid, commonly called phthalate esters, are a diverse group of compounds that have broad use in a wide array of industrial applications. Regulatory oversight of the manufacture, transport, use and disposal of phthalate esters has produced a large amount of data regarding the properties, environmental fate, exposure, and toxicity of these compounds. Such data are critical for the development of safe and accepted production practices, effluent discharge limits, and human exposure limits. The following chapters present, in detail, the information that has been collected regarding these properties. Keywords. Phthalate, Manufacture, Use, Releases, Regulation

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General Description

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Manufacture of Phthalate Esters . . . . . . . . . . . . . . . . . . .

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Nomenclature and Physical/Chemical Properties

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Chemical Interactions with Vinyl . . . . . . . . . . . . . . . . . .

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Uses of Phthalate Esters

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Disposal and Releases into the Environment . . . . . . . . . . . .

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Regulations and Phthalate Esters . . . . . . . . . . . . . . . . . .

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References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1 General Description Phthalate esters are widely used industrial chemicals. Higher molecular weight phthalate esters act as an additive which imparts flexibility in vinyl resins; this is the highest volume use of phthalates. Both linear and branched phthalate esters are used in the manufacture of vinyl articles. The linear esters provide superior low temperature properties to the finished vinyl products and also © Springer-Verlag Berlin Heidelberg 2003

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M.K. Stanley et al.

have lower volatility. The C8–C13 phthalate esters are the dominant vinyl plasticizers with di-2-ethylhexyl phthalate, diisononyl phthalate predominant and diisodecyl phthlatate. The lower molecular weight phthalates are used as plasticizers in some non-vinyl resins, including acrylics, urethanes and cellulosics. The various esters used in commerce have alkyl side chains containing from 1 to 13 carbon atoms. Table 1 contains a listing of the most common phthalate esters. Most of the high molecular weight phthalates esters are used in the manufacture of a wide variety vinyl goods, both commercial and consumer. The lower molecular weight phthalates, those with alkyl side chains from 1 to 4 carbon atoms, have a very broad use which includes consumer products and pharmaceuticals. This will be detailed in the use section.As plasticizers, phthalates are additives which improve the flexibility, processability and softness of vinyl. Phthalates with alkyl side chains lower than C6 are not often used alone as a plasticizer because of volatility concerns. In general, the factors that dictate selection of a phthalate or combination of phthalates for a particular application are functionality and economics of use [1]. Overall, phthalate esters are used because they combine qualities such as compatibility, permanence, efficiency and processability at reasonable cost. Compatibility problems with the vinyl resin preclude the use of phthalates esters of molecular weight higher than ditridecyl. In vinyl, dibutyl phthalate is only used in isolated cases in conjunction with higher molecular weight plasticizers to reduce volatility. Dipropyl and dipentyl phthalate, C3 and C5, are not available commercially in the United States.

2 Manufacture of Phthalate Esters The ortho phthalate esters are generally manufactured by the sequential addition of either branched or normal alcohols to phthalic anhydride in the presence of an acid catalyst. The alcohol manufacturing processes are stable, so although the phthalates produced from branched alcohols are complex substances, they are not variable. Phthalate esters are products of simple esterification reactions, which can be carried out readily in heated kettles with agitation and provision for water removal.While some plants produce phthalates by the batch method, newer, highly automated plants operate continuously, particularly if they emphasize a single product. The purity requirements for commercial plasticizers are very high and phthalate diesters are usually colorless and mostly odorless. The reaction usually requires an excess of alcohol, which is readily recycled. Diisononyl phthalate (DINP) is a complex substance assigned two different CAS numbers. CAS number 68515-48-0 is manufactured from polygas branched olefin that is converted to alcohol moieties consisting mainly of 3,4-, 4,6-, 3,6-, 3,5-, 4,5- and 5,6-dimethyl-1-heptanol. The CAS number 28553-12-0 is produced from dimerized n-butene that is converted primarily to methyl octanols and dimethyl heptanols. This CAS number also represents DINP which is produced from n-butene and isobutene that are converted to alcohols, with 60% consisting of methylethyl hexanols. The two types of DINP are considered commercially

Introduction

3

interchangeable. Other phthalates that are complex mixtures are diisodecyl phthalate (DIDP) and the linear phthalates D610P and D711P.

3 Nomenclature and Physical/Chemical Properties The physical and chemical properties of phthalate esters are well documented and will be discussed in detail in this handbook. A summary of some of their physical properties is contained in Table 1. The phthalate esters discussed here are liquids at room temperature. The diesters derived from the lower molecular weight alcohols such as DMP and DEP are colorless fluids of low viscosity, but phthalate esters become more viscous and oily as the size of the alkyl side-chain increases. They have low freezing points, many well below 0 °C (see Table 1). Generally, the water solubility of the alkyl phthalate ester varies inversely with the length of the alkyl side chain. DMP is the most hydrophilic and water soluble of the esters. The C10, C11 and C13 esters are the most hydrophobic and least water soluble (< 0.001 mg/l). Most of the dialkyl phthalates are soluble in common organic solvents such as benzene, toluene, xylene, diethyl ether, chloroform and petroleum ether [2]. In many cases alternate names are used in the literature and in commerce for the common phthalate esters. Table 1, while not intending to be exhaustive, lists these synonyms. Note the DOP (“dioctyl phthalate”) is used as a synonym for DEHP (di(2-ethylhexyl)phthlate).

4 Chemical Interactions with Vinyl Incorporating phthalate esters into a polymeric matrix reduces the glass transition temperature of the polymer [3]. Phthalate esters are not bound to the polymer with covalent chemical bonds and are therefore able to migrate to the surface of the polymer matrix where they may be lost by a variety of physical processes. Nevertheless, various chemical-physical attractive forces hold the phthalate ester tightly within the vinyl matrix, so that such migration occurs at a very low rate. Retention in the polymer matrix is one of the main factors in considering which phthalate ester to use. The ester must be sufficiently nonvolatile to remain in the compound during its mixing and formation stages

5 Uses of Phthalate Esters Uses of phthalate esters can be broadly split into three general categories – vinyl plasticizers, plasticizers for non-PVC polymers and other minor specialized applications. In the United States, DEHP, DINP and DIDP account for 52.2% of phthalates consumed. Linear phthalates account for 21.4% of the consumption and the 26.4% balance of US consumption includes all other phthalates esters [4].

Dimethyl Phthalate Diethyl Phthalate Di-n-Butyl Phthalate Diisobutyl Phthalate Butylbenzyl Phthalate Dihexyl Phthalate

Diisoheptyl Phthalate

Di-n-Octyl Phthalate Di (n-Hexyl, n-Octyl, n-Decyl) Phthalate Di(2-Ethylhexyl) Phthalate Diisononyl Phthalate

Diisodecyl Phthalate

Di(Heptyl, Nonyl, Undecyl) Phthalate

Diundecyl Phthalate Ditridecyl Phthalate

DMP DEP DnBP DIBP BBP DHP

DIHP

DnOP D610P

DIDP

D711P

DUP DTDP

DINP

DEHP

Phthalate Ester

Abbreviation

C30H50O4 C34H58O4

C26H42O4

C28H46O4

C26H42O4

C24H38O4

C24H38O4 C25H40O4

C22H34O4

C10H10O4 C12H14O4 C16H22O4 C16H22O4 C19H20O4 C20H30O4

Formula

Table 1. Physical properties of phthalate esters

28553-12-0 68515-48-0 26761-40-0 68515-49-1 3648-20-2 68515-44-6 68515-45-7 111381-89-6 111381-90-9 111381-91-0 3648-20-2 119-06-2 68515-47-9

131-11-3 84-66-2 84-74-2 84-69-5 85-68-7 84-75-3 68515-50-4 7188-89-6 6815-44-6 117-84-0 25724-58-7 68515-51-5 117-81-7

CAS No.

249-079-5 271-090-9 247-977-1 271-091-4 222-804-9 271-086-7 271-087-2 – – – 222-884-9 204-294-3 271-089-3

204-214-7 247-210-0 271-091-4 204-211-0

205-011-6 201-550-6 201-557-4 201-553-2 201-622-7 201-559-5 271-093-5 276-15-8

EINECS No.

447.7 {432.7–474.7} 530.8 {506.8–544.8}

418.6 {362.6–474.7}

446.7 {432.7–446.7}

418.6 {418.6–432.6}

390.6

390.6 404.6 {334–447}

363

194.2 222.2 278.4 278.4 312.4 334.4

Molecular Weight

–9 –37

4) octanol-water partition coefficients can be extracted with high recovery. Baltussen et al. [27] introduced magnetic stir bars coated with silicones as alternative solventless extraction method (stir bar sorptive extraction, SBSE). The stir bars are placed in 10–50 mL samples and are stirred on a magnetic stirrer for 30–120 min.After extraction, the stir bar is removed from the sample and placed in a thermal desorption unit for thermal desorption and on-line GC-MS analysis. The same methodology is used for air monitoring (see below). For semivolatile compounds, such as phthalates, a 10 mm-long stir bar coated with a 0.5 mm film of dimethylpolysiloxane (PDMS volume = 24 µL) is used. The amount of coating is 50 times higher and consequently higher recoveries are obtained for compounds with lower Kow values. Stir bar sorptive extraction has been used for various environmental samples and was also used for the extraction of semi-volatile compounds, including phthalates, in beverages. The possibilities of stir bar sorptive extraction, followed by thermal desorption-GC-MS are demonstrated in Fig. 10.A 10 mL rainwater sample was collected directly in a 20 mL vial. The PDMS-coated stir bar was introduced and extraction was performed over 2 h. The resulting chromatogram (extracted ion chromatogram at m/z 149) shows the presence of DiBP (16.01 min), DBP (18.39 min) and DEHP (29.59 min). The concentrations were 4.60 ppb (DiBP), 0.53 ppb (DBP) and 0.25 ppb (DEHP). These data clearly illustrate that transport through air is an important pathway of distribution of phthalates in the environment. This analysis also shows that very high sensitivities can be obtained by SBSE. In this application, the mass spectrometer was operated in scan mode and the signal-to-noise ratio for DEHP is higher than 40. The detection limit is thus lower than 25 ppt (ng L–1) in scan mode and lower than 5 ppt in SIM mode.

29.59 min = DEHP)

Fig. 10. Extracted ion chromatogram of stir bar sorptive extraction of phthalates from rainwater (16.01 min = DiBP, 18.39 min = DBP,

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5 Analysis of Phthalates in Sediments, Soils and Sewage Sludges In comparison with water samples, the determination of phthalates in solid (or semi-solid) samples usually requires two steps: an extraction step and a clean-up step. Solid samples include soils, sediments, sludges and solid waste. Soil samples are normally collected as grab samples by using a stainless steel drill, which allows sampling at different depths. Typical concentration levels for phthalates in soil are in the order of 10–1000 µg kg–1 dry mass. Sampling of sediments is used for the study of historical phthalate contamination and for the determination of local contamination and biodegradation. Sampling techniques that can be used for sediment sampling are described by Parkman and Remberger [21] and by Braaten et al. [22]. Concentration levels of phthalates largely depend on the sampling site. The average concentrations are of the same order of magnitude as the soil samples. Sludge samples from wastewater treatment are important samples to monitor input sources and to study biodegradation. Moreover, determination of phthalates in treated (digested and dried) sludge may be important. In at least one European country (Denmark) there is a defined maximum allowable level for DEHP in sludge to be used as an agricultural fertiliser. Phthalates entering the wastewater treatment plant via the wastewater influent are in general well removed from the aqueous stream in part by biodegradation and in part by removal in the sludge. Typical phthalate concentrations in treated sewage sludge are between 10–100 mg kg-1 dry mass. Dried sludges usually have a 20–30% dry mass content and can be analysed in a similar way to soils and sediments. Wet sludges (dry mass < 5%) can be analysed as such or after concentration of the solids and removal of the aqueous phase (centrifugation, filtering). Solid samples can be stored at –20 °C during several weeks. 5.1 Extraction

For the extraction of phthalates from solid (or semi-solid) matrices, various techniques have been used. Soxhlet extraction, whereby an organic solvent is heated and the condensed vapours are percolated through the solid samples held in a filter cartridge (thimble), is still considered as the reference method for the extraction of semi-volatile pollutants from solid environmental samples. Soxhlet extraction is a slow extraction procedure (4–24 h) and uses relatively large amounts of solvent (100–250 mL per sample). Due to the large solvent consumption, contamination is therefore a major issue with Soxhlet extraction. Soxhlet extraction has been used by Steffen and Lach [28]. Sediment samples were first freeze-dried. Then 30–50 g samples were extracted with 200 mL toluene over 8 h. For some samples, sulfur compounds were removed prior to GC-MS analysis by sonication of the extract in combination with copper powder. During the past years, automated and miniaturised extraction techniques are slowly replacing classical Soxhlet extraction. These techniques include automated Soxhlet extraction (Soxtec), shaking, ultrasonic extraction, microwave assisted

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extraction (MASE), accelerated solvent extraction (ASE) and supercritical fluid extraction (SFE).Automated miniaturized Soxhlet extraction (Soxtec, Soxtherm) is faster than Soxhlet extraction and solvent consumption is reduced. Further reduction of solvent consumption is possible by using supercritical fluid extraction. In SFE, pressurized carbon dioxide is brought to a pressure and temperature above its critical pressure (75 bar) and critical temperature (35 °C), resulting in a supercritical fluid. A supercritical fluid can be considered as a dense gas, with gas-like viscosity and flow characteristics and liquid-like solvating characteristics. Supercritical fluid extraction has been used successfully for a broad range of applications. SFE was used by Kolb et al. [29] for the extraction of phthalates from sewage sludge. The extraction was performed at 60 °C with carbon dioxide modified with 5% hexane as the extraction solvent and a three-step pressure program (5 min at 200 bar, 5 min at 300 bar and 20 min at 400 bar). The extraction efficiencies were higher than 90% for DBP, BBzP and DEHP, and were 84% for DiNP and 79% for DiDP. The standard deviation was 2–7%. Currently, supercritical fluid extraction is replaced by accelerated solvent extraction (ASE). In ASE, similar equipment is used, but the supercritical fluid is replaced by a classical organic solvent (dichloromethane, hexane, etc.). The solvent is pumped into an extraction vessel containing the sample. The solvent in the thimble is pressurized (up to 5000 psi) and heated (typically at 100°C). After extraction, the solvent is eluted in a vial. The extraction is completely automated and in comparison to Soxhlet extraction, solvent consumption is drastically reduced. Accelerated solvent extraction was used by Lettinski et al. [25]. Typically 15 g soil sample is mixed with anhydrous sodium sulfate (or Hydromatrix). The sample is placed in an extraction thimble and extracted with dichloromethane at 2000 psi and 100 °C for15 min. Recoveries of phthalates were 80–100%. The main problem was the relatively high background values that were typically around 36 µg kg–1 for DiBP, 24 µg kg–1 for DBP and 35 µg kg–1 for DEHP. Probably one of the most simple and cheap extraction methods is shaking or sonication. Vikelsoe et al. [4] used a simple shaking method for the determination of phthalates in soil samples. The samples were not dried before extraction. After addition of internal standard, a 50 g sample was extracted with 100 mL dichloromethane by shaking for 4 h. The blank values were around 6 µg kg–1 for DEHP. Low solvent consumption in combination with relatively short extraction times are also obtained by ultrasonic extraction. Braaten et al. [22] used sonication for the determination of phthalates in sediments by using 5 g wet sample. Firstly, 2 mL acetonitrile was added and a 10 min ultrasonic extraction was performed. Then, 2 mL acetonitrile and 3 mL hexane were added and the ultrasonic extraction was repeated for 30 min. The organic phase was isolated, washed with water and the hexane phase was cleaned on aluminium oxide. A similar method was applied by Parkman and Remberger [21] to the determination of phthalates in Swedish and Dutch sediments. Liu [6] used a triple sonication extraction of 2 g sample (mixed with 20 g sodium sulfate) with each time 20 mL 1 : 1 dichloromethane : hexane for the determination of phthalates in marine sediments. Paxeus [30] also used ultrasonic extraction of dried sludge with MTBE or a 1 : 1 mixture of hexane and acetone. Recoveries were higher than 90% by

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using three 10 min extractions. The organic phase (upper layer) is removed after each extraction and replaced by fresh solvent. Zurmuhl [31] demonstrated that similar extraction recoveries are obtained by ultrasonic extraction in comparison to Soxhlet extraction. 5.2 Clean-Up Procedures

Since extracts of soils, sediments and sludges often contain other contaminants or co-extracted compounds such as plant sterols, clean-up is often needed. A simple clean-up method based on solid-phase extraction is described by Braaten et al. [22]. The extract from sediments are applied on a 500 mg neutral alumina column (activated at 400 °C, deactivated with 9% water). The column is rinsed with 3 mL hexane. The phthalate fraction is eluted with 3 mL of a 75% hexane/25% MTBE mixture. Liu [6] used 15 g alumina column (300 ¥10 mm i.d., alumina deactivated with 15% water) with 1–2 cm anhydrous sodium sulfate on top. After applying the sample, the column is eluted with 30 mL hexane (waste), then with 30 mL 10% dichloromethane in hexane (PCB fraction) and finally with 30 mL 50% dichloromethane in hexane. This last fraction contains the phthalates. In some cases, an additional clean-up on a 7.5 g Florisil column was used. The column was rinsed with 30 mL dichloromethane (waste) and with 30 mL 5% acetone in dichloromethane. The phthalates are present in this last fraction. The method blanks for a 2 g sediment sample were: 1.3 µg kg–1 DiBP, 28.4 µg kg–1 DBP and 24.9 µg kg–1 DEHP. The detection limits were thus determined by the method blanks for these compounds. The reported recoveries were higher than 70% for DMP, higher than 80% for DEP, DiNP and DiDP, and higher than 90% for DiBP, DBP, BBzP and DEHP. 5.3 Determination of Phthalates in Sewage Sludge

The determination of phthalates in a sludge sample is demonstrated in Fig. 11. The sludge was obtained from a wastewater treatment plant. The extraction was performed by ultrasonic extraction with acetone : hexane according to the procedure described by Paxeus [30]. In this sample, DiBP, DBP, BBzP and DEHP are clearly detected. At the end of the chromatogram, the “hump” corresponding to DiNP and DiDP is also detected. In this sample, the concentration of DEHP, the most abundant phthalate, was around 30 mg g–1 dry mass.

6 Analysis of Phthalates in Air 6.1 Introduction

Transport in the atmosphere is an important pathway for the distribution of phthalates in the environment [32, 33]. Most of the phthalates are adsorbed onto

Fig. 11. GC-MS chromatogram obtained for a wastewater treatment plant sludge extract

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particulate matter [34–36] and earlier methods used for monitoring phthalates in air often focused on the collection and analysis of dust samples [37].Although the vapour pressures of phthalates are relatively low [23], phthalates can still be regarded as semi-volatiles and are also present in the aerosol or vapour phase. Since phthalates are present at trace levels (ng–µg m–3 level), direct analysis of air samples is not possible and enrichment techniques are needed. Enrichment of compounds in air can be done by active or passive concentration of the solutes on adsorbents or sorbents. For the determination of phthalates adsorbed on particulates, active or passive sampling on filters is used. 6.2 Analysis of Total Phthalate Concentrations in Air

For the determination of total phthalate concentrations (gaseous + aerosols + particulates), sampling on a sorbent tube is used. Most widely used adsorbents are activated carbon, porous polymers like Tenax (2,6-diphenylphenylene oxide polymer) and resins (XAD-2) [38–47]. Recommended methods for the sampling and analysis of phthalates (OSHA CIM, OSHA 104, NIOSH 5020) also include trapping of the phthalates present as aerosols or in the vapour phase on a combination of a polyurethane foam (PUF) plug and a resin cartridge. Phthalates adsorbed to particulate material can also be collected first on a glass fibre filter. In these generic methods, typically 60–240 L of air are sampled at a rate of 1000 mL min–1. It should be noted that with these methods contamination problems can occur due to PVC filter holders or plasticized rings that are used to hold the glass fibre filters. These materials can contain phthalates and consequently lead to high blank values. After sampling, the phthalates are desorbed from the adsorbents by liquid extraction, or by thermal desorption. In the case of particulates concentrated on filters, liquid extraction is used. The extraction procedure for particulates on filter media is very similar to the extraction of phthalates from solid samples (soil, sediment). Ultrasonic extraction, Soxhlet extraction, accelerated solvent extraction and others can be used [41, 47]. The advantage of liquid desorption is that quantitative extraction of the solutes can easily be obtained, the extract can be further fractionated or purified and the final extract can be analysed several times. Liquid desorption, however, lacks sensitivity. Assuming a limit of detection of 10 pg per compound with mass spectroscopic detection, a concentration of minimum 10 ppb is required in the final extract. By using liquid extraction of the sorbent or filter, concentration to 1 mL and a 1 µL injection, at least 1000 L of air should be sampled to quantify 10 ng m–3. Otake et al. [46] used for instance 100 mg charcoal tubes and sampled for 72 h at 1 L min-1 (total sample 4.3 m3). After sampling, desorption was done with 1 mL toluene and the extract was analysed by GC-MS. A detection limit of 4 ng m–3 could be obtained. Thermal desorption, on the other hand, has the advantage of higher sensitivity because all sorbed compounds can be quantitatively transferred to the GC and the detector. The same sensitivity (10 pg in detector) can thus be obtained by sampling only 1 L of air. The limiting factor in thermal desorption is the desorption efficiency. The adsorption of high-molecular weight compounds is strong and a high energy

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(high temperature) is needed for quantitative desorption especially when classical adsorbents are used (e.g. carbon-based materials, Tenax and Porapak). Tenax adsorption traps can be used successfully for the low-molecular weight phthalates (typically DEP, DiBP, DBP), but for the high-molecular weight isomeric mixtures (DiNP, DiDP) poor recoveries have been observed. Pre-concentration by sorptive enrichment with silicones offers a useful alternative and was introduced by Baltussen et al. [48, 49]. Air was sampled through a tube packed with a bed of 100% polydimethylsiloxane (PDMS). The polymeric material is above its glass transition temperature (Tg) at sampling temperatures (0–30 °C) and the solutes are, in contrast to classical sampling systems, sorbed into (dissolved in) the liquid stationary PDMS phase rather than adsorbed onto an active surface. The sampling traps allow sufficient enrichment at a relative high sampling speed and quantitative thermal desorption can be performed at moderate temperatures. Moreover, the material is highly inert and has a high thermal stability. However, for the enrichment of phthalates, it was observed that 100% PDMS traps resulted in too many bleeding compounds (low-molecular weight silicones) that interfered in the extracted ion chromatograms at m/z 149 [50]. This is the most abundant and the quantification ion for phthalate determination. The dimethylsiloxane oligomers give m/z 73 and m/z 147 as most abundant ions but because of the silicium isotopes the ion m/z 149 is also observed in the spectra.An alternative method was found by coating a silicone layer on an inert support in a 5% (w/w) concentration of 5%. Thermal desorption tubes (4 mm i.d. ¥ 180 mm) were filled with 100 mg of the material and were conditioned at 300 °C for 2 h. The ‘blank’ profile showed only some traces of volatile silicon fragments, but these did not interfere with the phthalate peaks. By using these traps, ng m–3 concentrations of phthalates can be measured. Since pre-concentration on PDMS tubes is based on sorptive enrichment, the breakthrough volumes (Vb) of phthalates can be calculated directly from the theory developed by Lövkist and Jönsson [51]. The values for some important phthalates are summarised in Table 6 and vary between 17 L for dimethyl phthalate up to 75 ¥ 106 L for diisodecyl phthalate when sampling is performed at a rate of 500 mL min–1. If the sample volume is limited to 15 L, all phthalates are quantitatively trapped. Assuming a mass spectrometric detection limit of 10 pg, the sensitivity of the thermal desorption-GC-MS method is 10 pg/15 L or 0.7 ng m–3. In practice, a Table 6. Breakthrough volumes (L) of several phthalates when

sampling is performed on PDMS tubes (100 mg 5% PDMS) at a flow rate of 500 mL min–1 at room temperature (20 °C) Compound

Breakthrough volume (L)

DMP DEP DIBP DBP DEHP DIDP

17 23 13 ¥ 102 19 ¥ 102 17 ¥ 106 75 ¥ 106

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limit of quantitation of 3 ng m–3 was obtained due to typical background levels of DiBP, DBP and DEHP around 20–30 pg. A typical sampling and analysis procedure based on sorptive enrichment and thermal desorption-GC-MS is summarized as follows [50]. Samples are aspirated through a thermal desorption tube containing 100 mg 5% PDMS sorbent by using a universal air sampling pump (SKC, Dorset, UK) at a nominal flow of 500 mL min–1 for 30 min (sample volume = 15 L). After sampling, the sampling tubes are closed and stored in an airtight container or wrapped in aluminium foil. After sampling, the tubes are placed in a thermal desorption unit (TDS-A, Gerstel GmbH, Muelheim, Germany), mounted on an Agilent 6890 GC coupled to an Agilent 5973 mass selective detector. Desorption is started by programming the tube to 300 °C. The released solutes are transferred from the sampling tube through a heated fused silica capillary into a cryotrap (PTV inlet). During thermal desorption, the PTV is cooled to –100 °C and the phthalates are trapped in an empty liner. After 10 min, the thermal desorption is completed and the solutes are injected into the capillary column by rapidly heating the PTV to 300 °C at 10 °C s–1. The chromatographic and mass spectroscopic conditions are the same as described above (Table 3). As for other sample matrices, a potential problem in the trace analysis of phthalates in air is contamination. It is very important that the contact between stored sample tubes and laboratory air is minimised, since DiBP and DBP were found to be two important contaminants originating from the surrounding lab atmosphere. A typical chromatogram of the analysis of laboratory indoor air is presented in Fig. 12. The measured concentrations of DiBP and DBP are 200–700 ng m–3. If tubes are left unprotected in the laboratory, they will adsorb phthalates and this leads to overestimated data. The limit of detection (LOD) of the method with a sampling speed of 500 mL min–1 and a sampling time of 30 min (15 L sample) is of the order of 1 ng m–3 for the single isomer phthalates and around 10 ng m–3 for the mixed isomer phthalates. For DiBP, DBP and DEHP, background values under optimised (clean) conditions can range between 2–3 ng m–3 and values below these limits are questionable for these compounds. The sorptive enrichment-thermal desorption-GCMS method was validated in the concentration range 3 to 3000 ng m–3. The correlation coefficients of the linearity curves were higher than 0.9970 for the single isomer phthalates DMP, DEP, DiBP, DBP, BBzP and DEHP, and 0.9786 for DiNP and 0.9782 for DiDP. The relative standard deviation of five replicate analyses at a level of 100 ng m–3 was between 2.3% (for DEP) and 9.2% (for DEHP). As a typical example of phthalate monitoring in air, the concentration of phthalates was measured in a greenhouse and in the surrounding atmosphere. Outside samples were taken at 1, 10 and 100 m distances. The resulting data are summarized in Fig. 13. Inside the greenhouse, the concentrations of the detected phthalates were respectively: 226 ng m–3 DiBP, 156 ng m–3 DBP, 48 ng m–3 BBzP and 309 ng m–3 DEHP. At 100 m distance, only background levels of DiBP, DBP and DEHP were measured and the concentrations decrease with increasing distance from the greenhouse. It should also be noted that dynamic (active) sampling by using either adsorbents or sorbents measures total phthalate concentrations, since particulates will

Fig. 12. Phthalate of laboratory air at m/z 149 on a 5% PDMS tube. Peaks 1 DEP, 2 DiBP, 3 DBP, 4 DEHP

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Fig. 13. Phthalate concentrations (ng m–3) in air samples in and outside a greenhouse

also be trapped on the material that acts as a filter. If a glass filter is placed before the (ad)sorbent tube, the particulates can be monitored separately. However, the differentiation between particle-bound (and adsorbed) phthalates and the phthalates in gaseous and aerosol phase by such a sampling train (filter+sorbent) is questionable. It can be expected that phthalates initially present in the gaseous phase can also be concentrated on a filter or on dust collected on the filter during sampling due to adsorption (so the concentration in the sorption tube is too low). In contrast, phthalates initially adsorbed on particles and trapped on a glass fibre filter can be purged out towards the sorption tube (giving too high values for the “gaseous+aerosol fraction”). This effect has already been clearly demonstrated by Schulz and Püttmann [37]. With a 1 h sampling time, measured phthalate concentrations were higher than the values obtained by using a 24 h sampling. This effect was called the “blowing off ” problem. Therefore, we recommend the use of adsorbents or sorbents for the determination of total phthalate concentrations. 6.3 Passive Sorptive Sampling of Phthalates in Air

Passive sampling is a very suitable technique for the measurement of air quality in indoor and working environment [52]. For passive sampling, a sorbent is put in a holder. This may be placed in a specific place in the room or used as a personal sampler. Sampling is performed over several hours or days and concentrations are expressed as time weight averages (TWA). The (ad)sorbed solutes are desorbed with a suitable solvent and the extract is analysed. Recently, solid-phase micro-extraction (SPME) with a PDMS coating immobilized on a fused silica

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fibre, was introduced as an alternative sampling device for passive diffusion sampling [53–55]. After sampling, the fibre is introduced in a hot GC inlet and the concentrated compounds are thermally desorbed in the GC. The major constraint of the SPME method is the relatively small amount of sorbent that is present on the fibre (0.5 µL). For this reason, the recently developed PDMS-coated stir bars can be an interesting alternative. It has been demonstrated that these stir bars can be used to sample volatiles in the headspace of liquid or solid samples [56]. In passive sampling with sorptive extraction, the PDMS polymer is exposed to an air sample during a relative long period (hours, days) and the pollutants present at an initial concentration CS0 are enriched until equilibrium is reached between the gas and the PDMS polymer phase. This distribution can be defined for a given component at a given temperature as the distribution coefficient KPDMS/air : • CPDMS KPDMS/S = 9 (1) CS• • where CPDMS and CS• are the solute concentrations in the PDMS phase and air sample at equilibrium state, respectively. Since passive sampling is usually performed in open areas, where the sample volume is much larger than the volume of the PDMS phase, the concentration CS• at equilibrium does not significantly differ from the initial concentration CS0. This last value can thus easily be calculated: n (2) or n = VPDMS · CS0 · KPDMS/S (3) CS0 = 968 VPDMS · KPDMS/S

where n the absolute amount of solute that is enriched in the PDMS polymer and VPDMS is the volume of the PDMS phase. Equation (3) shows that n is directly proportional to the amount of sorbent VPDMS . This implies that if a coated bar containing 50 µL PDMS is used instead of an SPME fibre (0.5 µL PDMS) the sensitivity is increased by a factor 100. Passive sampling with a PDMS-coated bar was applied to the determination of phthalates in a room with PVC flooring. A clean PDMS-coated bar containing 50 µL PDMS was exposed to the indoor environment for 24 h.After sampling, the phthalates were analysed by thermal desorption-GC-MS. The resulting extracted ion chromatogram at m/z 149 is shown in Fig. 14. The high abundance of DiBP, DBP and DEHP illustrates the extremely high enrichment of the phthalates in the PDMS polymer phase. Since phthalates adsorbed on particulates are not enriched by this sampling method, passive sampling with sorptive extraction allows differentiation between phthalates that are present in the vapour or aerosol phase and phthalates adsorbed onto particulates. Only the phthalates in vapour or aerosol phase are measured. Therefore, passive sorptive sampling is complementary to dynamic sorptive enrichment. 6.4 Dust Analysis

An important part of the human phthalate intake from the atmosphere may be attributed to the inhalation of dust particles on which plasticizers are adsorbed.

Fig. 14. Passive diffusion extraction (PSSE, 50 µL PDMS) of phthalates in a carpeted house

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Hence, attention has been paid to the determination of phthalates in dust. The hazardous impact of phthalates adsorbed on dust particles depends on the particle size of the dust.According to the American Conference of Governmental Industrial Hygienists (ACGIH) [57] and European regulation [58], the inhalable dust fraction is defined as the fraction of particulates with a median particle size smaller than 100 µm. The thoraric fraction is defined as the fraction of particulates with a median particle size smaller than 10 µm. The respirable fraction is defined as the fraction of particulates with a median particle size smaller than 4 µm. Inhalable dust is deposited in the respiratory trajectory (mouth and gullet), while respirable particles are deposited anywhere in the gas-exchange region, including the lung bladders. Thoraric dust is retained before the lung area. Dust sampling can be done by using vacuum cleaners, glass fibre filters in combination with high-volume air sampling pumps or with special dust samplers. A detailed description of dust sampling is given in VDI method 4300 [59]. Special dust samplers allow differentiation between the total inhalable and respirable dust filtration [60]. By using these samplers, the total inhalable dust fraction is aspirated at a flow of about 2 L min–1 onto a glass fibre filter (pore size 1.0 µm) through a broad cassette inlet. This sampler discriminates larger particulate matter. Smaller respirable particles can be collected with a cyclone-type sampler. The recent models are constructed of plasticizer-free plastics that eliminate electrostatic problems. This avoids repellence of the particles and contributes to high sampling efficiencies. In both cases, sampling is typically performed over several hours. Total dust concentrations are measured gravimetrically. Phthalates adsorbed on the dust fraction can be determined after liquid extraction of the filter. However, in our experience, it is extremely difficult to obtain reliable data on phthalates on the dust fraction collected in these special samplers. More reliable data are obtained by sampling larger amounts of dust by highvolume sampling, followed by dust fractionation by sieving and phthalate determination by liquid extraction and GC-MS. Fractionation of dust samples can be done by using stainless steel sieves or by centrifugation. Extraction of phthalates from dust can be performed by the methods described for solid samples [47, 61]. Alternatively, thermal extraction can be used, especially if limited amounts of sample are available. A few mg of sample can be placed in a thermal desorption tube and the tube is analysed in the same way as for air samples. An example is given in Fig. 15, showing the thermal extraction-GC-MS analysis of a house dust sample. The measured phthalate concentrations were 90 mg kg–1 DBP, 200 mg kg–1 DiHP, 700 mg kg–1 DEHP and 300 mg kg–1 DiNP. This sample clearly shows the presence of the C7-isomeric mixture of diisoheptyl phthalate eluting before DEHP.

Fig. 15. Thermal extraction-GC-MS analysis of house dust

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7 Analysis of Phthalates in Biota (Vegetation, Milk, Fish) 7.1 Introduction

The determination of phthalates in biological matrices such as vegetation, fish or milk is more difficult due to the complexity of the matrix. In general, the methodology for the determination of phthalates in biota can be divided into two classes, depending on the fat content of the matrix. For samples with a relatively low fat content (< 1%), phthalates can be extracted by using the same methods as described for soils, sediment and other solid samples. During the extraction, other constituents such as sterols, pigments, flavanoids, waxes, fatty acids, etc will be co-extracted. After extraction, a cleanup method is needed. As clean-up methods, column chromatography or solidphase extraction can be used. In these clean-up methods, the separation of phthalates and co-extracted compounds is based on differences in polarity. For fatty matrices, the main problem is the co-extraction of fats. Since lipids have a polarity similar to the polarity of phthalates, it is difficult to remove them with methods based on column chromatography or solid-phase extraction. The fat matrix can be removed by a clean-up method based on size exclusion chromatography (gel permeation chromatography, GPC). In size exclusion chromatography, compounds are separated according to the molecular volume (ª molecular mass). Larger molecules cannot enter the pores of the GPC packing material and elute before smaller molecules that can enter the small pores. GPC was first described for the fractionation of pesticides from lipid matrices [62–63]. However, this liquid chromatography method is more and more considered as a very valuable sample preparation technique for all determinations of organic contaminants in lipid matrices. On GPC columns, phthalates with molecular weights in the range 200–400 Da are separated from lipids (MW around 800 Da). Classical GPC separations are performed on large (40 cm ¥ 25 mm i.d.) low-pressure columns, operated at 5 mL min–1 [62–63]. The method can be miniaturized and solvent consumption can be largely reduced by using HPLC equipment and high-pressure GPC columns. Automated GPC clean-up can be performed with a system consisting of an isocratic HPLC pump, an autosampler allowing the injection of 500 µL fat extract, a temperature-controlled column oven, a variable UV detector (optional) and a fraction collector. The separation is performed on small-bore columns (e.g. 300 mm ¥ 7.5 mm i.d. PL-Gel with 5 µm particles and 5 nm pore size), operated at 1 mL min–1 dichloromethane. The 5 nm pore size is important, since it allows the separation of organic compounds in the 100–1000 mass range. Best resolution is obtained on two columns in series (total length = 60 cm). Solvent consumption is approximately 5–10 times lower in comparison to the classical GPC method. The phthalate fraction (3–5 mL) is collected automatically.After collection, the fraction is concentrated and this clean-up step can eventually be followed by an additional clean-up method according to polarity by using column chromatography or SPE.

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7.2 Analytical Procedure for the Determination of Phthalates in Vegetation

Vegetation samples normally do not contain high levels of lipids. For the determination of phthalates in plant material, first a liquid-solid extraction is used.All techniques described for solid samples (Soxhlet, shaking, ultrasonic extraction, ASE, SFE) can also be used for the extraction of phthalates from plant material. High recoveries are obtained by using a simple shaking or ultrasonic extraction method. These methods are also fast, cheap and relatively small amounts of solvents are used. For the extraction, two approaches can be used. In the first approach [6], vegetation samples are first homogenised with a blender. Approximately 5 g (wet) sample is spiked with internal standard(s) and mixed with 30 g anhydrous sodium sulfate (pre-baked at 450 °C) in a mortar until a dry powder is obtained. The dried sample is extracted with 30 mL of a 1 : 1 mixture of hexane and dichloromethane by sonication for 10 min. After the suspended particles are settled, the supernatant is removed. The extraction is repeated twice with fresh solvent and finally the three organic fractions are combined and concentrated to 1 mL under nitrogen. In the second approach [7], the sample is not dried with sodium sulfate. Approximately 10 g homogenised (wet) sample is weighed in a 40 mL I-Chem vial. After addition of the internal standard(s) and 10 mL acetone, the sample is extracted for 15 min by sonication. Then 10 mL cyclohexane is added and the sonication extraction is repeated for another 15 min. After this first extraction, the vials are placed on a shaking machine for 30 min. Finally, the vials are again placed in the ultrasonic bath for 15 min. After completion, the extraction vial is centrifuged. An aliquot (5 mL) of the (upper) cyclohexane phase is transferred to another tube and concentrated to 1 mL Clean-up of the extracts can be done by column chromatography or by solidphase extraction. The procedures are identical to the procedures described for solid samples [6,22]. By using column chromatography, the concentrated extract is transferred onto an alumina column packed with 15 g deactivated alumina (15% water w/w) and with a 1–2 cm bed of anhydrous sodium sulfate on top. The column is eluted with 30 mL hexane, followed by 30 mL 10% dichloromethane in hexane and finally 30 mL 50% dichloromethane in hexane. This last fraction contains the phthalates. This fraction is concentrated to 1 mL and analysed. The clean-up can be miniaturized by using a 500 mg alumina cartridge. After applying the extract, the cartridge is washed with 3 mL hexane and the phthalates are eluted with 3 mL 75% hexane-25% MTBE. The limit of detection of these methods is in the order of 2–10 µg kg–1 wet sample for the single isomer phthalates (and 20–100 µg kg–1 wet sample for the mixed isomer phthalates). However, method blanks are typically in the order of 10–20 ppb, especially for DiBP, DBP and DEHP. For these compounds, the limit of quantitation is around 20–40 µg kg–1 wet sample (set to two times the background level).

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7.3 Analytical Procedure for the Determination of Phthalates in Milk or Edible Oils and Fat

For the determination of phthalate in milk samples the following method was developed [7]. Milk samples are first homogenised by shaking. Milk powders are dissolved in water (1 g/10 mL). Approximately 5 g milk sample is then extracted with 20 mL of a 1 : 1 cyclohexane/acetone mixture in a 40 mL I-Chem vial. The vials are shaken on a shaking machine for 30 min.After completion, the vials are centrifuged and the supernatant is transferred to a pre-weighed I-Chem vial. The solvent is evaporated under nitrogen and the fat content is measured. The residue is dissolved in dichloromethane and internal standard (d4-DEHP) is added. The amounts of solvent and internal standard are adjusted to give approximately 50 mg fat and 100 ng internal standard per mL dichloromethane. The solution was then homogenised in a vortex agitator. The separation of the fat from the phthalate containing fractions is then performed by gel permeation chromatography (GPC). For the determination of phthalates in edible oils and fat, the oil or fat sample can by diluted directly in dichloromethane to a concentration of 50 mg fat per mL. If the sample solution is not clear, some water might be present and this can be removed by adding some anhydrous sodium sulfate to the sample. The clear solution is also fractionated by GPC. For the GPC separation of the dichloromethane solution, 500 µL is injected onto a two column combination. This combination consisted of a 5 cm ¥ 7.5 mm i.d. PL-Gel pre-column and two 30 cm ¥ 7.5 mm i.d. ¥ 5 µm PL-Gel 5 nm columns. The mobile phase is dichloromethane at 1 mL min–1 flow rate. UV detection at 220 nm is used to monitor the effluent. Phthalates typically elute in a window between 20 and 23 min (3 mL fraction). This fraction is automatically collected. The total run time is 30 min. To the collected fraction, 100 µL cyclohexane is added and the extract is concentrated to 100 µL. This extract is analysed by GC-MS. The limit of detection of this method is around 20 ng g–1 fat. DiBP, DBP and DEHP are detected in the method blanks, but the levels are constant around 20–50 ng g–1 fat. Hence, the limit of quantification for these phthalates is 40–100 ng g–1 fat (two times background level). For milk samples with 3% fat content, this corresponds to a quantification limit of 1–3 ng g–1 milk. The recovery of the GPC clean-up method was tested by spiking an olive oil sample at a 500 ppb level with DMP, DEP, DiBP, DBP, BBzP and DEHP and at a 5 ppm level with DiNP and DiDP. The olive oil was tested before and no phthalates were detected at concentrations above 50 ppb (500 ppb for DiNP and DiDP). The results from the non-spiked oil were therefore considered as the “procedure blanks”. D4-DEHP (internal standard) was added to the dichloromethane solution. The sample was analysed in triplicate. The linearity was tested by spiking an olive oil sample at seven levels (100, 250, 500, 1000, 2500, 5000 and 7500 ng g–1 fat) (¥ 10 for mixed isomers). The mean recovery, standard deviation (RSD %) and linearity (correlation coefficient r2) are listed in Table 7. In general, good recoveries (>80%) are obtained, except for butylbenzyl phthalate for which the recovery is lower. This is probably due to a slightly different behaviour of this

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Recovery (%) RSD (%) Linearity (r2)

DMP

DEP

DiBP

DBP

BBzP

DEHP

DiNP

DiDP

103 15.7 0.9956

134 15.7 0.9946

104 9.5 0.9992

138 11.8 0.9962

69 16.0 0.9947

127 12.1 0.9978

85 11.6 0.9841

81 10.6 0.9903

compound in sample clean-up (GPC). The correlation coefficients are better than 0.99 for all phthalates, except for DiNP (0.98). This correlation shows that the method can be used in the concentration range from 100–7500 ppb for the single isomer phthalates and from 1–75 ppm for the mixed isomer phthalates. An alternative and automated method for the determination of phthalates in oil and fat matrices was described by Pacciarelli et al. [64]. In this method, an online HPLC-GC method was used. The sample fractionation was performed by HPLC in straight (normal) phase mode. Fractionation of the triglycerides was done on a silica column (100 ¥ 4.6 mm i.d. Lichrosorb SI-100, 5 µm) using 1 : 1 dichloromethane/cyclohexane (with 0.5% acetonitrile) as mobile phase at 1 mL min–1. The fraction containing DEHP was automatically transferred in the GC and the triglyceride matrix is eluted afterwards. 7.4 Analytical Procedure for the Determination of Phthalates in Fish

Depending on the type of fish and available sample size, the fish samples can be analysed as whole fish, as fillet (muscle with skin removed) or the analysis can be performed on specific organ samples (liver, stomach). For diet studies, the fillet sample represents the edible part of the fish. For contamination studies, whole fish samples or organ analyses give relevant information. Due to their lipophilic character, phthalates accumulate in the fat. The analytical methods for the determination of phthalates in fish consist in general of a fat (+phthalate) extraction step, followed by a clean-up step. Different extraction techniques can be used. These include ultrasonic extraction, Soxhlet extraction,ASE, etc. For cleanup, gel permeation chromatography and column chromatography or solid phase extraction can be used. A fish matrix is quite complex and it is advisable to use both GPC plus an additional alumina column clean-up based on polarity differences to remove fats and other interfering compounds that are co-extracted. Liu et al. [6] used ultrasonic extraction. The samples are homogenised unfrozen. Analytical subsamples, 5 g wet weight, are spiked with d4-labelled standards and mixed with 30 g anhydrous sodium sulfate in a mortar until a dry powder is obtained. This homogenised and dried sample is extracted with 30 mL dichloromethane by sonication for 10 min and shaking for another 10 min. After the suspended particles are settled, the supernatant is removed and the extraction is repeated twice with fresh solvent. Finally, the three dichloromethane fractions are combined and concentrated to 1 mL. The extracts are first cleaned by GPC. The phthalate fraction is concentrated and then transferred onto an alu-

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mina column packed with 15 g deactivated alumina (15% water w/w) and with 1–2 cm bed of anhydrous sodium sulfate on top. The column is eluted with 30 mL hexane, 30 mL 10% dichloromethane in hexane and finally 30 mL 50% dichloromethane in hexane. This last fraction contains the phthalates. This fraction is concentrated to 1 mL and analysed. By using this method, Liu et al. [6] obtained detection limits around 2 ng g–1. Due to method blanks, the limit of quantification for DBP was around 20 ng g–1 and around 40 ng g–1 for DEHP. The recovery was 70–95% for the single isomer phthalates, measured by GC-MS and 75–110% for the mixed isomer phthalates measured by LC-MS. The same preparation procedure was used by David et al. [7]. In this procedure, 5 g homogenised fish sample was mixed with internal standard (d4-DEHP) and with 15 g pre-cleaned sodium sulfate. Extraction was performed by sonication with 20 mL cyclohexane for 15 min. The extract was centrifuged and the supernatant was removed. The extraction was performed another time with fresh solvent and the combined extracts were concentrated under nitrogen and fractionation by GPC, using the method described for milk samples. An example of the GPC separation for a fish extract is given in Fig. 16. The chromatogram obtained for a DEHP reference standard is overlayed. The DEHP peak elutes on the tail of the lipid peak. The fraction between 15 and 17 min is collected, concentrated and analysed by GC-MS. In this fraction, some sterols elute that cannot be separated from DEHP. These compounds do not interfere with the further analysis. The GC-MS chromatograms are shown in Fig. 17. The extracted ion chromatogram at m/z 149 shows the presence of DEHP. The extracted ion chromatogram at m/z 153 shows the presence of the internal standard (d4-DEHP). In this case, no additional clean-up by column chromatography or by SPE was used. The recovery for DEHP measured at three spike levels between 2 ng g–1 (wet sample) and 40 ng g–1 (wet sample) were 89–93%. The relative standard deviation at 40 ng g-1 (wet sample) was 9.7%. Letinski [25] used accelerated solvent extraction for the extraction of phthalates from fish tissue.Approximately 1 g fish sample was mixed with sodium sulfate and extracted with a 90% hexane/10% ethyl acetate mixture at 120 °C and 1600 psi. This extract was purified on alumina. The limit of quantification was 150 ng g–1 fish sample and recovery was 98% for the recovery matrix spike.

8 Sample Preparation Methods for Phthalic Acid Mono-Esters Phthalic acid monoesters are more polar than phthalates and therefore the sample preparation methods must be adapted. For the determination of monoesters in blood, plasma and urine samples, several methods are described in the literature [8–10, 12, 13, 17, 18]. For the determination of monoesters in environmental samples, only limited data are available. Suzuki et al. [14] used solid-phase extraction on polymeric phase disks (styrene divinylbenzene) for the determination of monoesters in riverwater samples. The waters were acidified to pH 2 before extraction to ensure the monoesters are in the protonated form (not ionised). Extraction efficiency was higher than 72% (except for monomethyl phthalate).

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Fig. 16. GPC fractionation of phthalates in fish extract

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After extraction, the monoesters must be derivatised prior to GC analysis. Alternatively, HPLC of non-derivatised monoesters can be performed (see above). Recently, a method has been developed for the analysis of phthalic acid monoesters in water samples by using an in situ derivatisation, followed by stir bar sorptive extraction and thermal desorption GC-MS [19]. One mL of water sample is placed in a 20 mL glass vial and spiked with internal standard solution. 500 µL of a 2 : 1 (v/v) mixture of ethanol and pyridine is added and the

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Fig. 17. GC-MS chromatograms of DEHP and internal standard (d4-DEHP) extracted from fish

mixture was homogenised by vortex agitation. After addition of 100 µL of ethyl chloroformate, the mixture was again vortex homogenised and placed in an ultrasonic bath for 15 min. During vortex and ultrasonic agitation gas evolved from the vial. The mixture was finally diluted with 10 mL water. The derivatised monoesters are extracted by stir bar sorptive extraction (SBSE), followed by thermal desorption-GC-MS. The limit of detection (LOD) is in the order of 0.05 µg L–1 for MBP and MEHP. The analysis of a sample spiked at 1 µg L–1 levels is shown in Fig. 18. The monoesters MBP, MEHP, MiNP and MiDP are easily detected.

Fig. 18. Determination of phthalic acid monoesters in water by in situ derivatisation and SBSE-thermal desorption-GC-MS

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9 Conclusions During the past years, various sample preparation and analytical methods have been developed for the determination of phthalic acid diesters in different environmental samples. The major problem in phthalate analysis is the risk of contamination. Precautions have to be taken into account to minimize this risk and to control the background values. Based on the authors’ personal experience, the following methods are recommended for the different matrices. Clean water samples can be extracted by methods based on solid-phase extraction. For contaminated water samples, miniaturized liquid-liquid extraction can be used. Sediments, sludges and soil samples can be extracted by using ultrasonic extraction. Biota samples can also be extracted by sonication. The extracts are purified by gel permeation chromatography, followed by column chromatography or solid-phase extraction. The analysis of the single isomer phthalates can be done by GC-MS. HPLC-MS offers an interesting alternative for the analysis of mixed isomer phthalates. For the analysis of monoesters in environmental samples, new methods are currently under development. The extraction and clean-up method used for phthalates need to be adapted. Specifically, the monoesters must be derivatised prior to GC-MS analysis. Alternatively, they can be analysed without derivatisation by using LC-MS.

10 References 1. US EPA methods 525 (drinking water), 606 and 625 (waste water) and 8060 and 8270 (solid waste). US Environmental Protection Agency, Cincinnati, Ohio 45268, USA 2. ISO draft international standard 18856, ISO. Geneva, Switzerland 3. Vikelsoe J, Thomsen M, Johansen E (1998) Sources of phthalates and nonylphenols in municipal waste water, technical report no 225 and 268. National Environmental Research Institute (NERI), Roskilde, Denmark 4. Vikelsoe J, Thomsen M, Johansen E, Carlsen L (1999) Phthalates and nonylphenols in soil, technical report no 268. National Environmental Research Institute (NERI), Roskilde, Denmark 5. George C, Prest H (2001) Agilent technologies application note Nr 5988–2244EN 6. Lin, Zhong-Ping, Ikonomou MG, Hongwu J, Macintosh C, Gobas FAPC (2002) Environ Sci Technol submitted for publication 7. David F, Tienpont B, Sandra T, Sandra P (2002) submitted for publication 8. Sjöberg P, Bondesson U (1985) J Chromatogr 344:167 9. Sjöberg P, Bondesson U, Sedin E, Gustafsson J (1985) Eur J Clin Invest (1985) 15:430 10. Sjöberg P, Bondesson U, Sedin E, Gustafsson J (1985) Transfusion 25:424 11. Marschall H-M, Green G, Egestad B, Sjövall J (1988) J Chromatogr 452:459 12. Egestad B, Green G, Sjöberg P, Klasson-Wehler E, Gustafsson J (1996) J Chromatogr B 677:99 13. Haam D, Vandenbroek P, Jongeneelen F (1993) Int Arch Occupat Environ Health 64:555 14. Suzuki T, Yaguchi K, Suzuki S, Suga T (2001) Environ Sci Technol 35:3757 15. Tienpont B, David F, Sandra P (2002) submitted for publication 16. Barry Y, Labow R, Keon W, Tocchi M, Rock G (1989) J Thorac Cardiovasc Surg 97, 900 17. Shintani H (2000) Chromatographia 52:721 18. Blount B, Milgram K, Silva M, Malek N, Reidy J, Needham L, Brock J (2000) Anal Chem 72:4127

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19. Tienpont B, David F, Sandra P (2002) submitted for publication 20. Furtmann K (1994) Fresenius J Anal Chem 348:291 21. Parkman H, Remberger M (1995) Phthalates in Swedish sediments, IVL Report Publ no 1167, p 21+appendix 22. Braaten B, Berge JA, Berglind L, Baekken T (1996) Occurence of Phthalates and Organotins in sediments and water in Norway. Norwegian Institute for Water Research (NIVA) report SNO 3552–96, p 45 23. Cousins I, Mackay D (2000) Chemosphere 41:1389 24. Van der Velde E, Korte de G, Versteegh A (1998) Determination of phthalate esters in water by SPE or in-vial extraction with GC-MS analysis: how to avoid the contamination problem. Paper presented at 20th international symposium on capillary chromatography, Riva del Garda, Italy, 26–29 May 1998 25. Letinski DJ (1999) Lecture presented at Workshop Nov 4–5 26. Pawliszyn J (1999) Applications of solid phase micro-extraction, RSC chromatography monographs. Royal Society of Chemistry, Letchworth, UK (ISBN 0–85404–525–2) 27. Baltussen E, Sandra P, David F, Cramers C (1999) J Microcolumn Sep 11:737 28. Steffen D, Lach G (2000) Phthalate und Trichlosan in Sedimenten und Schwebstoffen Niedersächsischen Gewässer, Niedersächsisches Landesamt für Ökologie, report 10/2000) 29. Kolb M, Welte K, Mettenleiter S, Trinkmann A (1997) Wasser Boden 49:57 30. Paxéus N (1999) submitted for publication 31. Zurmühl T (1990) Analyst 115:1171 32. Thomas GH (1973) Environ Health Perspect 3:23 33. Giam CS, Chanm HS, Neff GS, Atlas EL (1978) Science 199:419 34. Cautreels W, van Cauwenberghe K (1978) Atm Environ 12:1133 35. Cautreels W, van Cauwenberghe K, Guzman LA (1993) Sci Total Environ 8:47 36. Thúren A, Larsson P (1990) Environ Sci Technol 24:554 37. Schulz H-M, Püttmann W (1993) Analysis of saturated hydrocarbons, fatty acids and phthalic acid esters in air particulate matter of a city area (Aachen). Wissenschaft und Umwelt 2/1993, p 131 38. Chang LW, Atlas E, Giam CS (1985) Int J Environ Anal Chem 19:145 39. Figge K, Rabel W, Wieck A (1987) Fresenius J Anal Chem 327:261 40. Vainiotalo S, Pfaffli P (1990) Ann Occup Hyg 34:585 41. California Environmental Protection Agency (1992) Monitoring of phthalates and PAHs in indoor and outdoor air samples in riverside, California. Contract no A933-144, Dec 1992 42. Fisher J, Ventura K, Prokes B, Jandera P (1993) Chromatographia 37:47 43. Fujimoto T, Takeda N, Taira T, Iikawa R (1995) Kurin Tekunoroji 5:45 44. Bartulewicz J, Bartulewicz E, Gawlowski J, Niedzielski J (1996) J Chem Anal (Warsaw) 41:753 45. Weiling G, Xiku W (1997) Huanjing Huaxue 16:382 46. Otake T, Yoshinaga J, Yanagisawa Y (2001) Environ Sci Technol 35:3099 47. Rudel RA, Brody JG, Sprengler JD, Vallarino J, Geno PW, Sun G, Yau A (2001) J Air Waste Manage Assoc 51:499 48. Baltussen E, Janssen HG, Sandra P, Cramers CA (1997) J High Resol Chromatogr 20:385 49. Baltussen E, David F, Sandra P, Janssen HG, Cramers CA (1998) J High Resol Chromatogr 21:332 50. Tienpont B, David F, Sandra P, Vanwalleghem F (2000) J Microcolumn Sep 12:194 51. Lövkist P, Jönsson JA (1987) Anal Chem 59:818 52. Namiesnik J, Gorecki T (2000) LC-GC Europe, September 2000, pp 678 53. Martos PA, Pawliszyn J (1997) Anal Chem 69:206 54. Martos PA, Pawliszyn J (1999) Anal Chem 71:1513 55. Khaled A, Pawliszyn J (2000) J Chromatogr 892:455 56. David F, Sandra P (2001) Passive sorptive sampling for indoor air monitoring using polydimethylsiloxane coated stir bas, paper 740, The Pittsburgh conference on analytical chemistry and applied spectroscopy, 4–9 March 2001

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57. ACGIH technical committee on air sampling procedures (1984) Particle size-selection sampling in the workplace. ACGIH, Cincinnati, Ohio 58. EN 481 (1993) European committee for standardisation CEN 1993-07–27 59. VDI 4300 (1999) Measurement of indoor air pollution, sampling of house dust. Verein Deutscher Ingenieure, Düsseldorf 60. Mark D, Vincent JH (1986) Ann Occup Hyg 30:89 61. Bruns-Weller E, Pfordt J (2000) Z Umweltchem Okotox 12:125 62. Specht W, Tilkes M (1980) Frezenius Z Anal Chem 301:300 63. Thier H-P, Zeumer H (1987) Manual of pesticide residue analysis, vol 1. VCH, Weinheim, p 75 64. Pacciarelli B, Müller E, Schneider R, Grob K, Steiner W, Fröhlich D (1988) J High Resol Chromatogr 11:135

The Handbook of Environmental Chemistry Vol. 3, Part Q (2003): 57– 84 DOI 10.1007/b11463

Physical-Chemical Properties and Evaluative Fate Modelling of Phthalate Esters Ian T. Cousins 1 · Donald Mackay 1 · Thomas F. Parkerton 2 1 2

Canadian Environmental Modelling Centre, Environmental and Resource Studies, Trent University, Peterborough, Ontario, K9J 7B8, Canada. E-mail: [email protected] Exxon Mobil Biomedical Sci. Inc., Hermeslaan 2, 1831, Machelen, Belgium

A review is presented of the physical-chemical properties and reactivity of the phthalate esters including a discussion of how these properties control their partitioning and fate in the environment. The air and water solubilities decrease by orders of magnitude from the short alkyl chain phthalates such as dimethyl phthalate (DMP) to the long alkyl chain phthalates such as di-2-ethylhexyl phthalate (DEHP). The octanol-water partition coefficient, which is a measure of hydrophobicity, increases by orders of magnitude with increasing alkyl chain length and this increase is mainly controlled by the reduction in water solubility rather than an increase in octanol solubility. This increase in hydrophobicity results in strong sorption of the higher molecular weight phthalates to organic matter.Air-water partition coefficients (or Henry’s law constants) also increase with increasing alkyl chain length. However, the greater evaporative potential of higher molecular weight phthalate esters from water is offset by sorption to suspended matter in surface waters. Phthalates have high values of KOA suggesting that they will be appreciably sorbed to aerosol particles, soil and vegetation. From available data obtained under environmental conditions, half-lives of phthalates in environmental media are proposed. Systematic differences in reactivity or half-life are apparent, with the primary biodegradation half-life tending to increase with increasing alkyl chain length. In contrast, the opposite pattern is observed for the air oxidation half-life. A series of evaluative modelling calculations is described to illustrate how the physical-chemical properties result in differences in environmental partitioning behaviour, persistence and transport potential. In comparison to other organic chemical classes, model results indicate that phthalates are not environmentally persistent or subjected to significant long-range transport. Although the overall environmental persistence of the higher molecular weight phthalates tends to increase, KOA and thus the propensity to partition to aerosols, vegetation and soils also increases, thereby reducing the potential for long-range transport. Recommendations for future research on physical-chemical properties of phthalate esters for environmental fate assessment are discussed. Keywords. Phthalate ester, Structure, Physical-chemical property, Model, Fate

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Introduction

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Structure-Property Analysis of Physical-Chemical Properties

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Observations on Physical-Chemical Properties

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Physical State . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64 Solubility in Water . . . . . . . . . . . . . . . . . . . . . . . . . . 64 Vapour Pressure . . . . . . . . . . . . . . . . . . . . . . . . . . . 66

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© Springer-Verlag Berlin Heidelberg 2003

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3.4 3.5 3.6 3.7

Air-Water Partition Coefficient . . Octanol-Water Partition Coefficient Octanol-Air Partition Coefficient . Data Gaps . . . . . . . . . . . . . .

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Degrading Reactions . . . . . . . . . . . . . . . . . . . . . . . . . 72

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Evaluative Fate Modelling with the EQC Model . . . . . . . . . . . 74

5.1 5.2 5.3 5.4

EQC Level I Modelling . . . . . . . . . . . . . . . . . . . . EQC Level II Modelling . . . . . . . . . . . . . . . . . . . . EQC Level III Modelling . . . . . . . . . . . . . . . . . . . Estimating Persistence and Long-Range Transport Potential with the TaPL3 Model . . . . . . . . . . . . . . . . . . . .

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

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References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82

1 Introduction Phthalate esters are widely used as plasticizers, serving as important additives that impart flexibility to polymers including poly(vinyl chloride) (PVC), polyvinylacetates, cellulosics and polyurethanes [1]. The stability, fluidity and low volatility of high-molecular mass phthalate esters make them ideal for use as plasticizers. The variety of possible chemical structures of phthalate esters results in a wide range of physical-chemical properties and hence environmental partitioning behaviour for this class of compounds. This wide range of properties is principally a result of the variation in the length of the alkyl chains substituted on the diester groups. The names, molecular formulae, molar masses and melting points of 22 phthalate esters are listed in Table 1. The objectives of this chapter are to review the published physical-chemical and reactivity data for the phthalate esters, seek relationships between chemical structure and properties and determine how these properties will influence partitioning between abiotic media in the environment with the use of evaluative environmental fate models. The accumulation of phthalate esters in biotic media (i.e. food webs) is the focus of a separate chapter in this volume.

2 Structure-Property Analysis of Physical-Chemical Properties Physical-chemical properties which can be measured readily in the laboratory with a view to determining environmental partitioning include: solubility in water, vapour pressure, the Henry’s law constant (H), the octanol-water partition coefficient (KOW) and the octanol-air partition coefficient (KOA). There are few direct measurements of Henry’s law constants for the phthalate esters and no

Physical-Chemical Properties and Evaluative Fate Modelling of Phthalate Esters

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Table 1. List of phthalate esters studied and their associated molar masses, molar volumes and melting points

Phthalate ester

Abbreviation

Molar mass (g mol–1)

Le Bas molar volume (cm3 mol–1)

Dimethyl phthalate Diethyl phthalate Diallyl phthalate Dipropyl phthalate Di-n-butyl phthalate Disiobutyl phthalate Di-n-propyl phthalate Butylbenzyl phthalate Diisohexyl phthalate Di-n-heptyl phthalate Di-n-octyl phthalate Butyl 2-ethylhexyl phthalate Di(n-hexyl, n-octyl, n-decyl) phthalate a Di(2-ethylhexyl) phthalate Diisooctyl phthalate Di-n-nonyl phthalate Diisononyl phthalate Di-n-decyl phthalate Diisodecyl phthalate Di(heptyl, nonyl, undecyl) phthalate a Diundecyl phthalate Ditridecyl phthalate

DMP DEP DAP DPP DnBP DIBP DnPP BBP DHP DIHpP DnOP BOP 610P DEHP DIOP DnNP DINP DnDP DIDP D711P DUP DTDP

194.2 222.2 246.2 250.3 278.4 278.4 250.3 312.4 334.4 362.5 390.6 334.4 404.6 390.6 390.6 418.6 418.6 446.7 446.7 418.7 447.7 530.8

206.4 254.0 283.6 298.4 342.8 342.8 387.2 364.8 431.6 476.0 520.4 416.6 542.6 520.4 520.4 564.8 564.8 609.2 609.2 564.8 653.6 742.4

a

Melting point (°C) 5.5 –40 – – –35 –58 – –35 –27.5 – – –37 –4 –46 –46 – –48 – –46 100 days). BBP is a special case in that it does not contain two straight alkyl chains in its structure and thus the mechanism for primary degradation is likely to be different. Measured data for BBP from Staples et al. [4] suggest that its aerobic biodegradation half-lives in natural waters, soils and sediments are 0.4 day to 8 ¥ 104 days, 9.6 days and 1.6–2.2 days, respectively. The value of 8 ¥ 104 days seems to be erroneous because there are five other data points in the range 0.4–1.4 days. There is only one data point for aerobic biodegradation in soil for BBP, which is particularly disappointing because soil is the primary medium of accumulation for BBP. Degradation rates of phthalate esters in anaerobic media are slower, but the models used in this chapter only treat aerobic environmental media. Only surface soils (top 5 cm) and sediments (top 3 cm) are treated. The above analysis of measured biodegradation half-life data from Staples et al. [4] has been used to allocate approximate half-lives by using a semi-decade logarithmic scale for water, soil and sediment compartments in the EQC Level II and III simulations (Table 5). This approximate allocation takes account of the large uncertainty in measured biodegradation half-lives. We have taken a conservative approach in our allocation of half-lives and assigned half-lives that are near to the top of the range reported by Staples et al. [4]. This conservative approach results in estimated half-lives that are higher than those suggested in the chapter of this handbook focussing on environmental degradation rates of phthalates. However, it is believed that a conservative approach is appropriate for allocation of half-lives because degradation studies are often conducted at a constant 25 °C, whereas the environment is often at a lower temperature, some studies use inoculums and some allow the microbial population to become acclimated. Furthermore, some microcosm studies may not separate losses from degradation from losses due to partitioning to sediments and volatilisation.

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I.T. Cousins, D. Mackay and T.F. Parkerton

Table 5. Allocation of half-lives for use in Level II and III EQC simulations

Phthalate ester

DMP

DEP

DnBP

BBP

DEHP

Assumed reaction half-lives (h)

Class Mean Range Class Mean Range Class Mean Range Class Mean Range Class Mean Range

Air

Water

Soil

Sediment

5 550 300–1000 4 170 100–300 3 55 30–100 2 17 10–30 2 17 10–30

4 170 100–300 4 170 100–300 4 170 100–300 3 55 30–100 5 550 300–1000

5 550 300–1000 5 550 300–1000 6 1700 1000–3000 6 1700 1000–3000 7 5500 3000–10,000

6 1700 1000–3000 6 1700 1000–3000 7 5500 3000–10,000 6 1700 1000–3000 7 5500 3000–10,000

5 Evaluative Fate Modelling with the EQC Model Conducting evaluative assessments can provide invaluable insights into the characteristics of chemical behaviour in the environment. Because the environment considered is purely evaluative or hypothetical, there is no possibility of validation, but the equations used to describe partitioning, transport and transformation are identical to those used successfully in validated models of chemical fate in more defined environments. The aim is to establish the general features of chemical behaviour, namely, into which media the chemical will tend to partition, the primary loss mechanisms, the tendency for intermedia transport, the tendency to bioaccumulate, the tendency to undergo long-range transport and environmental persistence. Multimedia models of this type are widely used by the scientific community as useful tools for providing information on chemical fate and have also found acceptance in regulatory practice in a number of countries. The Equilibrium Criterion or EQC model, the model of choice here, has been described fully elsewhere [65]. Briefly, this model in the form of a computer program, deduces the fate of a chemical in Level I, II and III evaluative environments by using principles described by Mackay [36]. The EQC evaluative environment is an area of 100,000 km2 that is regarded as being representative of a jurisdictional region such as the US state of Ohio, or the country of Greece. EQC can simulate the chemical fate of a variety of different chemical class types, classified according to the data requirements to run a model simulation. Phthalate esters partition to all environmental media and are thus classified as type 1 chemicals

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Physical-Chemical Properties and Evaluative Fate Modelling of Phthalate Esters

for which all partition coefficients and Z values (fugacity capacities) must be defined [36]. 5.1 EQC Level I Modelling

EQC Level I modelling has been performed for the 22 phthalate esters listed in Table 1 for which physical-chemical properties have previously been estimated (Table 3). Level I EQC model results indicate that under equilibrium, steady state conditions, with no reaction, the vast majority of phthalates released will reside in soil, sediment or water with over 99% being distributed to these three media (Table 6). The low vapour pressures ensure that only small percentages partition to air. Phthalate esters with alkyl chains containing greater than five carbons partition almost exclusively to the organic carbon component of soil and sediment, whereas those with short alkyl chains (