Certification of butyltins and phenyltins in marine sediment certified reference material by species-specific isotope-dilution mass spectrometric analysis using synthesized 118Sn-enriched organotin compounds

A new marine sediment certified reference material, NMIJ CRM 7306-a, for butyltin and phenyltin analysis has been prepared and certified by the National Metrological Institute of Japan at the National Institute of Advanced Industrial Science and Technology (NMIJ/AIST). Candidate sediment material was collected at a bay near industrial activity in Japan. After air-drying, sieving, and mixing the material was sterilized with γ-ray irradiation. The material was re-mixed and packaged into 250 glass bottles (15 g each) and these were stored in a freezer at −30 °C. Certification was performed by use of three different types of species-specific isotope-dilution mass spectrometry (SSID–MS)—SSID–GC–ICP–MS, SSID–GC–MS, and SSID–LC–ICP–MS, with 118Sn-enriched organotin compounds synthesized from 118Sn-enriched metal used as a spike. The 118Sn-enriched mono-butyltin (MBT), dibutyltin (DBT), and tributyltin (TBT) were synthesized as a mixture whereas the 118Sn-enriched di-phenyltin (DPhT) and triphenyltin (TPhT) were synthesized individually. Four different extraction methods, mechanical shaking, ultrasonic, microwave-assisted, and pressurized liquid extraction, were adopted to avoid possible analytical bias caused by non-quantitative extraction and degradation or inter-conversion of analytes in sample preparations. Tropolone was used as chelating agent in all the extraction methods. Certified values are given for TBT 44±3 μg kg−1 as Sn, DBT 51 ± 2 μg kg−1 as Sn, MBT 67 ± 3 μg kg−1 as Sn, TPhT 6.9 ± 1.2 μg kg−1 as Sn, and DPhT 3.4 ± 1.2 μg kg−1 as Sn. These levels are lower than in other sediment CRMs currently available for analysis of organotin compounds.
Introduction
Organotin compounds have wide ranging chemical and toxicological properties. They are widely applied as stabilizers of plastics, as fungicides and pesticides, and as marine antifoulants [1]. Use of tributyltin (TBT) and triphenyltin (TPhT) as marine antifouling agents has led to their almost global dispersal. Although use of TBT and TPhT has been controlled in Japan since 1989, the compounds are still prominent in the coastal sea waters of Japan. Thus, these organotins and their degradation products, di- and mono-organotins, in sediment are frequently monitored to evaluate organotin pollution in water environment. Quantification of organotin species in environmental samples is very difficult, because of their instability and the low concentrations present. Because long-term and worldwide monitoring is necessary for these compounds, comparability of the analytical results is required [2].
The National Metrology Institute of Japan (NMIJ) has already developed, in 2001, a marine sediment certified reference material (NMIJ CRM 7301-a) for analysis of mono, di, and tributyltins (MBT, DBT, and TBT) [3]. Certification was performed using two different types of species-specific isotope-dilution mass spectrometric (SSID–MS) method—gas chromatography coupled with inductively coupled plasma mass spectrometry (SSID–GC–ICP–MS) [4] and SSID–GC–MS combined with a microwave-assisted extraction—in which a mixture of 118Sn-enriched butyltins synthesized in our laboratory was used as a spike. In 2005 we developed a new marine sediment certified reference material, NMIJ CRM 7306-a, certified for di and triphenyltins (DPhT and TPhT) and the butyltins by using three different types of SSID–MS method. In the SSID methods, 118Sn-enriched TPhT and DPhT newly synthesized from 118Sn-enriched metal, and a mixture of 118Sn-enriched butyltins previously synthesized from 118Sn-enriched metal [4] were used as spikes. The certification strategy for NMIJ CRM 7306-a was almost the same as that for NMIJ CRM7301-a [3] but with two improvements to ensure the reliability of the CRM. One was the sterilization of the material with γ-ray irradiation. NMIJ CRM7301-a was not irradiation-sterilized because of possible degradation of the organotin compounds. Yang et al. reported that significant losses of butyltins in methanol occurs during γ-ray irradiation whereas such degradation is minimal in a sediment matrix [5]. In our preliminary test, no substantial loss of organotins was observed on γ-ray irradiation, and thus irradiation-sterilization was performed for CRM7306-a to ensure long term stability. The other improvement was that a total of six combinations of four extraction methods and three SSID–MS methods were used for analysis of the organotin compounds to ensure the reliability of the certified values. The overview of the analysis for certification is shown in Fig. 1. Although SSID–MS methodology can correct most of the systematic errors that occur in the analysis, it can not compensate for non-quantitative extraction from the sample or for species re-conformation in the sample preparation process [6–11]. Indeed, quantitative extraction of MBT is not an easy task, because MBT is strongly adsorbed by sediment matrices, because of its polarity [12–14]. It has been reported that DBT is degraded during microwave-assisted extraction [6] and pressurized liquid extraction [7] when high microwave energy or high extraction temperatures are used, although species interconversion can be taken into account in SSID–MS techniques that furnish data for more than one enriched isotope, as has been described by Encinar and co-workers [6–9]. In this certification, therefore, the four different types of extraction method, mechanical shaking, ultrasonic, microwave-assisted, and pressurized-liquid extraction were adopted to avoid possible analytical biases that caused by non-quantitative extraction and degradation or interconversion of analytes. SSID–liquid chromatography (LC)–ICP–MS, was adopted as an SSID–MS method in addition to SSID–GC–ICP–MS and SSID–GC–MS, because organotin separation by LC does not need a derivatization step. Thus, use of SSID–LC–ICP–MS can avoid potential degradation or interconversion of analytes in the derivatization step. This paper mainly describes SSID–MS analysis for certification of NMIJ CRM 7306-a Marine Sediment for organotin compounds.
Fig. 1Overview of analysis for certification
Experimental
Preparation of sediment material
The starting material for the CRM was collected in a bay near industrial activity in the Kyushu area, Japan. Approximately 300 kg surface sediment was sampled and the water was removed by filtration. The sediment material was air-dried for 2 weeks at room temperature (25–28 °C). After removal of visible external materials (rock, shell, etc.) the sediment material was ground in a high-purity alumina ball-mill. Sediment powder that passed through a 104-μm sieve was homogenized using a pan-type mixer. The sediment powder was then bottled in glass bottles (60 g each) and irradiation sterilized (60Co, 20 kGy). The irradiated powder was re-mixed and homogenized by use of a rocking mixer RM-10S (Aichi Electric, Japan) as a precaution against an inhomogeneous degradation of organotin compounds during sterilization. Finally, the powder was placed in amber glass bottles (15 g each) and stored at −30 °C.
Conversion to dry mass basis
The concentrations of the constituents of this CRM are given on a dry-mass basis. A dry mass correction factor for sample humidity was determined by drying the sample at 110 °C. After 5 h the sediment sample reached constant weight, so it was decided the drying time would be 6 h in this experiment. The dry mass correction factor at the time of certification was 0.959±0.003 (average±standard deviation for ten different bottles).
Chemicals
Tributyltin (TBT) chloride, dibutyltin (DBT) chloride, and triphenyltin chloride (TPhT) were purchased from Wako Pure Chemicals (Osaka, Japan). Monobutyltin (MBT) chloride and diphenyltin chloride (DPhT) were purchased from Aldrich (Milwaukee, WI, USA). Tripropyltin (TPrT) chloride was purchased from Merck (Darmstadt, Germany). Standard solutions of each organotin compound except DPhT were separately prepared by dissolving each compound in methanol (pesticide analysis grade, Wako) to avoid any disproportionation reactions with other organotins. The standard solution of DPhT was prepared by dissolving DBTCl2 in 0.005 mol L−1 HCl in methanol, to prevent self-disproportionation. All standard solutions were stored at −20 °C and diluted working solutions were prepared daily before the analysis. Sodium tetraethylborate (NaBEt4) was purchased from Wako Pure Chemicals. A 5% m/v solution of NaBEt4 was prepared in a glove box that was purged with N2 gas. Other chemicals used were of analytical reagent grade. Pure water prepared by use of a Milli-Q water-purification system (resistivity 18 MΩ cm, Nihon Millipore Kogyo, Tokyo, Japan) was used throughout the experiments.
Synthesis of the 118Sn-enriched organotin compounds
The mixture of 118Sn-enriched butyltin compounds used as the spike for the ID methods were synthesized from 118Sn-enriched tin metal (92% enriched) purchased from Trace Science International (Ontario, Canada). The synthetic procedure has been described previously [4]. 118Sn-enriched DPhT and TPhT were individually synthesized from 118Sn-enriched tin metal (98% enriched) purchased from Nippon Sanso (Tokyo, Japan). The synthetic procedures were almost the same as those described elsewhere [15]. A mixture of ca. 0.5 g 118Sn metal and ca. 2 g iodine was introduced into a 50-mL round-bottomed flask containing acetic acid (10 mL) and acetic anhydride (10 mL). A small crystal of potassium iodide was added as catalyst and the mixture was gently heated to reflux. After cooling in an ice bath, orange crystals of 118SnI4 (1.6 g) were collected. 118SnI4 was placed in a 50-mLround-bottomed flask containing 30 mL diethyl ether and 1 mol L−1 phenyl magnesium bromide in THF was added dropwise. After heating to reflux for 3 h the white-pink precipitate was collected by filtration after the Grignard reagent had been hydrolyzed with water. The solid was dissolved in dichloromethane and the solution was filtered to remove insoluble impurities. The dichloromethane was removed to leave 118Sn-tetraphenyltin (TePhT) as a white solid; this was washed with ethanol. 118SnI4 (0.25 g) and 118Sn-TePhT (0.5 g) were mixed in a glass tube and heated to 200 °C. The reaction products were dissolved in ethanol and the insoluble products were removed by filtration. By the addition of 20% potassium fluoride solution, 118Sn-TPhT fluoride was obtained as an insoluble salt. The fluoride was isolated by filtration, washed with a minimum amount of ethanol, and then treated with conc. hydrochloric acid and extracted with pentane. The extracts were dried with MgSO4 and the pentane was removed to yield 118Sn-enriched TPhT chloride as a white powder. 118Sn-enriched DPhT dichloride was prepared by treating 18Sn-enriched TePhT with HCl. The product was extracted with pentane. The extracts were dried with MgSO4 and the pentane was removed to yield 118Sn-enriched DPhT dichloride.
Extraction procedure
Ultrasonic extraction
The ultrasonic extraction procedure for GC–ICP–MS and GC–MS was as follows. The sediment sample (ca. 0.5 g) was placed in a PFA centrifuge tube and spiked with an appropriate amount of the spikes. Then 2 g NaCl, 12 mL toluene containing 0.1% tropolone, and 10 mL acetic acid–methanol (1:1) were added to the tubes. The resulting mixtures were extracted in an ultrasonic bath for 30 min at 60 °C. After addition of 10 mL water the tubes were again shaken, for good phase separation, and then centrifuged at 3000 rpm for 5 min. Finally, the upper toluene layer was collected as the extract.
For LC–ICP–MS, the extraction solvent was replaced with 10 mL acetic acid–methanol (1:1) containing 0.1% tropolone, and the same extraction procedure was performed.
Mechanical shaking extraction
The sediment sample (ca. 0.5 g) was placed in a PFA centrifuge tube and spiked with an appropriate amount of the spikes. Then 2 g NaCl, 12 mL toluene containing 0.1% tropolone, and 10 mL 0.5 mol L−1 HCl in methanol were added to the tubes, and the resulting mixtures were mechanically shaken for 60 min. After addition of 10 mL water the tubes were again shaken, for good phase separation, and then were centrifuged at 3000 rpm for 5 min. Finally, the upper toluene layer was collected as the extract.
Microwave-assisted extraction
Closed-vessel microwave-assisted extraction (MAE) was performed. The microwave system used was Mars X (CEM, USA). The sediment sample (ca. 0.5 g) was placed in a PFA vessel and spiked with an appropriate amount of the spikes. Then 2 g NaCl, 12 mL toluene containing 0.1% tropolone, and 10 mL 1 mol L−1 acetic acid in methanol were added to the vessel. The microwave irradiation program was ramp to 120 °C in 10 min then hold for 10 min. The mixture was transferred to a PFA tube containing 10 mL water and the tube was shaken, for good phase separation, and then centrifuged at 3000 rpm for 5 min. Finally, the upper toluene layer was collected as the extract.
Pressurized liquid extraction
Pressurized liquid extraction (PLE) was performed with an ASE 200 instrument (Dionex, USA). The sediment sample (ca. 1 g) was placed in an 11-mL stainless extraction cell containing a glass filter and quartz sand and was spiked with an appropriate amount of the spikes. After filling the extraction vessel with quartz sand, the cell was placed in the PLE system. The extraction solvent used was 0.5 mol L−1 acetic acid in methanol containing 0.2% tropolone. The extraction conditions were: oven temp. 110 °C, pressure 10.3 MPa, static time and cycle was 5 min×4 times. The extract obtained was transferred to a PFA tube containing 10 mL water and 12 mL toluene, and the tube was shaken, for good phase separation, and then centrifuged at 3000 rpm for 5 min. Finally, the upper toluene layer was collected as the extract.
Derivatization procedure for GC–ICP–MS and GC–MS
The extracted solutions were transferred to PFA centrifuge tubes and 25 mL ammonium-acetate buffer (0.5 mol L−1, pH 5) and 0.2 mL 5% NaBEt4 solution were added. The tubes were mechanically shaken for 20 min, for ethylation and extraction, and then centrifuged to achieve phase separation. The toluene layers were also transferred to the tubes and mixed with 2 g anhydrous sodium sulfate to remove the water.
Clean-up procedure
Clean-up on a Presep-C Florisil cartridge (Wako, Japan) was performed after the derivatization, except for LC–ICP–MS measurement. The toluene layer was evaporated to ca. 2 mL under a stream of N2 gas and loaded on to a cartridge previously conditioned with hexane. The eluent from the cartridge was collected in a 15-mL glass centrifuge tube. Hexane (6 mL) containing 5% (v/v) diethyl ether was then also loaded on the cartidge and the eluent was collected in the glass tube. Finally, the collected eluent was evaporated to 0.2 mL under a stream of N2 gas and used as the measurement sample solution.
For the LC–ICP–MS measurement, a clean-up procedure using Presep-C18 cartridge (Varian, Australia) was performed. The extract was loaded on to a cartridge previously conditioned with acetone. The eluent from the cartridge was collected in a 15-mL glass centrifuge tube, evaporated to 0.2 mL under a stream of N2 gas, and used as the measurement sample solution.
Determination of organotin compounds by ID–GC–ICP–MS
The GC–ICP–MS used was that the GC (Agilent 6890GC) was coupled with an ICP–MS (HP4500, Yokogawa Analytical Systems, Tokyo, Japan) by means of a manufactured transfer-line (Yokogawa Analytical Systems). The GC column was HP-1 ms (30 m×0.32 mm i.d., 0.32 μm film thickness). The measured masses were m/z 118 and 120. The operating conditions and procedures for GC–ICP–MS were similar to those described elsewhere [4].
The concentrations of organotin compounds were calculated by inserting each value into the ID equation Eq. (1), based on a double-ID method [16, 17]:
where Cx is the analyte concentration in the sample (mol g−1), mx the mass of sample (g) used for the sample-spike mixture, my the mass of spike solution (g) used for the sample-spike mixture,  the mass of spike solution (g) used for the standard-spike mixture, mz the mass of standard solution (g) used for standard-spike mixture; R is the 120Sn/118Sn ratio in the sample–spike mixed solution, R′ is the 120Sn/118Sn ratio in the standard–spike mixed solution, Rx is the 120Sn/118Sn ratio in the sample solution, Ry is the 120Sn/118Sn ratio from the spike solution, Rz is the 120Sn/118Sn ratio in the standard solution, w is the correction factor for dry mass; n is the number of replicate measurements, and k, Ky, and K′ are the mass discrimination correction factors for each isotope ratio, which were calculated from the area ratio of 120Sn/118Sn for TPrT in each chromatographic run [4]. P is the purity of each organotin chloride reagent used for preparing the standard solutions, D is the dilution factor for each organotin chloride in the gravimetric dilution of the standard solutions, Mw the molecular weight of each organotin chloride, B the procedure blank, and E the variation introduced by extraction to the analytical results.
Determination of organotins by ID–GC–MS
The GC–MS used was Agilent model 6890GC/5983MSD (Agilent Technologies, Wilmington, DE, USA) with an HP-5 ms column (30 m×0.32 mm i.d., 0.25 μm film thickness). The measured masses were m/z 231 and 233 for MBT, m/z 261 and 263 for DBT and TBT, m/z 301 and 303 for DPhT, and m/z 349 and 351 for TPhT. The other operating conditions and procedures for GC–ICP–MS analysis were similar to those described previously [3].
The concentrations of organotins were calculated by inserting each value into Eq. (2), below, in which it was assumed that the relationship between mass ratio and peak ratio was linear.
where Cx is the analyte concentration in the sample (mol g−1), mx the mass of sample (g) used for the sample-spike mixture, my the mass of spike solution (g) used for the sample–spike mixture, RWL the ratio of the mass in the spike solution (g) to the mass in the standard solution (lower level), RWH the ratio of mass in the spike solution (g) to the mass in the standard solution (higher level), R the measured abundance ratio in the sample–spike mixed solution, RL the measured abundance ratio in the standard–spike mixed solution (lower level), RH the measured abundance ratio in the standard–spike mixed solution (higher level), w the correction factor for dry mass, P the purity of each organotin chloride reagent used for the preparing the standard solutions, D the dilution factor for each organotin chloride in gravimetric dilution of the standard solutions, Mw the molecular weight of each organotin chloride, B the procedure blank, and E the variation introduced by the extraction to the analytical results. The mass bias correction was not performed for GC–MS measurements. In this experiment we prepared several sample–spike mixed solutions of different mass ratios and chose the solutions having values of RL and RH slightly lower and higher than the R value for calculation.
Determination of organotin compounds by ID–LC–ICP–MS
The LC–ICP–MS system used consisted of a PU-712i HPLC pump (Inert model; GL Science, Tokyo, Japan), a Nanospace SI-2 auto-injection sampler (Siseido, Tokyo, Japan), and an Agilent 7500c ICP–MS (Yokogawa Analytical Systems, Japan). The LC column used was a Mitysil RP-18 GP (150 mm×2.0 mm i.d. 3 μm). All tubing used was 1/16 in (0.13 mm i.d.) PEEK. The mobile phase was acetonitrile–water–acetic acid–tetraethylammonium chloride–tropolone 65:30:5:0.1:0.075 (%, v/v). The mobile phase flow rate was 200 μL min−1 and the sample volume injected was 5 μL. The sample-introduction device for ICP–MS was modified from the default settings as follows: a PFA μflow of 50 μL min−1 was used as a nebulizer, the double-path Scott type spray chamber was cooled to −5 °C, a 1.0 mm dimmer injector torch was used, and additional O2 (0.1 L min−1) gas was mixed with the argon axial gas flow via a T-adaptor. The LC column was directly connected via a 1/16 PEEK adaptor. The concentrations of organotins were calculated by use of Eq. (1).
Homogeneity study
The between-bottle homogeneity of the CRM was determined by analyzing sub-samples taken from ten bottles selected from the lot of 250 bottles. All the organotin compounds were determined by ID–GC–ICP–MS after ultrasonic extraction. Analysis of variance (ANOVA) of the data was performed and mean squares within group (MSwithin) and among group (MSamong) were calculated. Then standard deviations between bottles (sbb) were calculated by use of Eq. (3):
If the measurement method was insufficient repeatable, the effect of analytical variation on the standard deviation between units ubb was calculated and used to estimate the inhomogeneity [18]. The ubb was calculated by use of Eq. (4):
where denotes the degrees of freedom of MSwithin.
Results and discussion
Homogeneity study
In the homogeneity study, between-bottle inhomogeneity (sbb) was only observed for TPhT, so ubb was used as uncertainty-derived inhomogeneity for TPhT. A relatively large sbb value (7.1%) was obtained for DPhT, although this value was not very large compared with uncertainties from other sources, for example the uncertainty of analytical results and between-method variance, as is shown later. These results indicate that this material may be considered homogeneous for butyltin and phenyltin analysis.
Stability of organotins in this material
We have been monitoring the stability of the butyltin compounds in NMIJ CRM7301-a since 2001. All three butyltins were sufficiently stable during storage for five years at −30 °C even though CRM7301-a was not sterilized. The new CRM 7306-a contains almost the same butyltin concentrations as CRM7301-a, its sample composition was also similar, and it was sterilized with γ-ray-irradiation. Thus, the butyltins in CRM 7306-a will also be stable for at least five years. Both the phenyltin and butyltin compounds in BCR CRM646, which was pasteurized at 100 °C, have been shown to be stable when the material is stored at below −20 °C [19]. Therefore, we decided to store CRM7306-a at −30 °C and to assess the stability of the material by measuring each organotin by ID–GC–ICP–MS over a period of 1 year, and that for CRM7301-a.
Characterization of the synthesized 118Sn-enriched organotins
Characterization of the synthesized 118Sn-enriched organotins was performed by GC–ICP–MS after mixing of each, dilution with methanol, and ethylation with NaBEt4.
Figure 2 shows the GC–ICP–MS chromatograms obtained at m/z 118 and 120 for the synthesized 118Sn-enriched organotins mixture. The 118Sn-enriched DPhT and TPhT could be synthesized individually. The molar ratio of the 118Sn-MBT, 118Sn-DBT, and 118Sn-TBT in the synthesized mixture measured by GC–ICP–MS was approximately 7:10:5.
Fig. 2GC–ICP–MS chromatograms of 118Sn and 120Sn obtained for the mixture of 118Sn-enriched organotin compounds. The chromatogram of 118Sn was shifted 9 s and 7500 cps for clarity
Ten replicate measurements of the ethylated 118Sn-enriched organotin mixture by GC–ICP–MS were performed to obtain the 120Sn/118Sn ratio for each compound. The mass bias was corrected with the in-run correction method with TPrT [4]. The 120Sn/118Sn isotope ratios for 118Sn-butyltins were 0.055±0.001 (mean±standard uncertainty, n=10), with no significant differences. The 120Sn/118Sn isotope ratios for 118Sn-enriched DPhT and TPhT were 0.0020±0.0002 and 0.00020±0.0003, respectively.
Assay for the standard solution of the organotins
To obtain a standard solution with a well-defined concentration, an assay was performed to determine the purity of the natural abundance organotin chloride reagents used to prepare the standard solutions. Details of the assay have been described elsewhere [4]. In brief, the amounts of organic and inorganic tin species impurities in the organotin chloride reagents were estimated by GC–ICP–MS. The inorganic impurities, except for Sn, were checked by ICP–MS. The non-organotin organic impurities were also checked by GC–FID. The water content of the organotin chloride reagents were evaluated with a Karl–Fisher coulometric titrator (Model CA-05; Mitsubishi, Japan).
The main impurities in the organotin chloride reagents used are water and tin species. Small amounts of inorganic and organotin impurities were observed in both the ethylated MBT and ethylated DBT solutions, and significant amounts of several organotin impurities were observed in the ethylated TBT, ethylated TPhT, and ethylated DPhT solutions. Among the organotin impurities observed in each ethylated organotin solution, inorganic tin, MBT, DBT and tetrabutyltin (TeBT), MPhT, and DPhT were identified from their retention times in GC–ICP–MS measurement and isomers of TBT (iso-TBT) and dioctyldibutyltin (DOcDBT) were identified from the fragment-ion patterns obtained by GC–MS. Two organotin impurities that could not be identified from their retention time were also observed in the ethylated-MBT solution, but the amounts of those were small (<0.03%). The purities of the organotin chloride reagent obtained were 99.4±0.2% for MBT, 99.5±0.2% for DBT, 96.8±0.2% for TBT, 98.6±0.2% for DPhT, and 98.8±0.2% for TPhT.
The stability of the standard solution during certification is also important. Van et al. reported that DPhT in methanol was not stable because of its disproportionation reaction, although solutions of the butyltins and TPhT were found to be stable [20]. DPhT also reacted with MBT in mixed solutions in methanol. In contrast, Arnold et al. reported that DPhT in methanol containing 0.01 mol L−1 HCl was stable for six months [21]. Hence, the stability test for DPhT in methanol with and without 0.005 mol L−1 HCl was performed by GC–ICP–MS after storage at −20 °C for one month. Redistribution of DPhT to MPhT and TPhT was observed for the methanol solution, as has been described elsewhere [20]. In contrast, no degradation or redistribution of DPhT was observed for the methanol solution containing 0.005 mol L−1 HCl. Plazzogna et al. reported that the redistribution reaction between monomethyltrialkyltin (R3SnMe) and dimethyltin dichloride (Me2SnCl2) in methanol is occurred with dissociation of Me2SnCl2 [22]. They also reported that the redistribution reaction can be prevented by addition of chloride ion, as NaCl, which inhibits the dissociation of Me2SnCl2 in methanol. For DPhT in methanol the disproportionation reaction occurred with dissociation of DPhTCl2, and was prevented by addition of HCl, which inhibited the dissociation. Consequently, the standard solution of DPhT was prepared by dissolving DBTCl2 in methanol containing 0.005 mol L−1 HCl to prevent the self-disproportionation reaction.
Evaluation of the degradation of DBT and TPhT during extraction
Because SSID–MS methodology cannot compensate for non-quantitative extraction from the sample, aggressive conditions are required for solid–liquid extraction of strongly adsorbed species, for example MBT. Degradation or interconversion of species can occur under the aggressive extraction conditions [6–11]. Indeed, it has been reported that degradation of DBT occurs during microwave assisted extraction [6] and pressurized liquid extraction [7] when high-energy microwaves or high extraction temperatures are used. Degradation of TPhT during sample pretreatment has also been reported [11, 12]. In this certification, therefore, degradation of DBT and TPhT during extraction was checked using an 117Sn-enriched DBT solution obtained from the LGC, UK [15] and the single solution of the synthesized 118Sn-enriched TPhT. Both 117Sn-enriched DBT and 118Sn-enriched TPhT solutions were used to spike the sediment sample and then the four extraction methods were performed. After derivatization and clean-up the extract was analyzed by GC–ICP–MS. Because the 118Sn-enriched TPhT solution does not contain any phenyltins, degradation of TPhT in each extraction was evaluated from the difference between the natural abundance isotope ratio and the measured ratios of 120Sn/118Sn for DPhT. On the other hand, the 117Sn-enriched DBT solution contains small amounts (ca. 0.7%) of 117Sn-enriched MBT as an impurity. Therefore, the degradation of DBT in each extraction was evaluated from the difference between the natural abundance isotope ratios and the ratios of 118Sn/117Sn for MBT, corrected for 117Sn-enriched MBT. In this experiment, the measured 120Sn/118Sn ratios for DPhT obtained for all four extraction methods closely matched the natural abundance, and the results suggest that no significant degradation of TPhT to DPhT occurred during extraction. The degradation of DBT to MBT was also not observed in ultrasonic extraction and mechanical shaking extraction. Slight degradation of DBT to MBT was observed in microwave-assisted extraction MAE (0.8%) and in pressurized liquid extraction PLE (0.5%); these degradation levels were lower than those reported by Encinar [6, 7] (2–3% in both extractions under their optimized conditions). These differences could be because of different extraction conditions. The sample size used in our experiments was two to four times that used by Encinar, and the concentrations of acetic acid we used were lower. In addition, tropolone was used as a chelating reagent in all the four extraction methods. Formation of the tropolone-chelating complex might prevent degradation. The slight degradation of DBT observed would not significantly affect the analytical results obtained by use of MAE and PLE because the typical uncertainty of isotope-ratio measurement by GC–ICP–MS in this study was in the range 0.5 to 1%, and the relative standard deviations of the analytical results for DBT obtained by use of MAE and PLE were approximately 3%. As is shown later, the analytical results obtained by use of the four different extraction methods were in good agreement within the range of uncertainties. These results indicate that analytical biases caused by degradation or inter-conversion of analytes in the extraction would be negligible in this certification.
Analytical results obtained by each method
Determination of the organotin compounds in the candidate material by the six analytical methods was performed for the certification. Figures 3, 4 to 5 show the GC–ICP–MS, GC–MS, and LC–ICP–MS chromatograms obtained for the sample extract spiked with the 118Sn-enriched organotin compounds. It is apparent from Figs. 3 and 4 that adequate sensitivity for all the organotins was obtained in GC–ICP–MS and GC–MS measurements. Adequate sensitivity for the organotin compounds, except for DPhT, was also obtained in LC–ICP–MS; the sensitivity for DPhT was insufficient to obtain a reliable isotope-ratio measurement. The relative standard deviation for DPhT in LC–ICP–MS measurement was approximately 10% in triplicate measurement, and thus variation of the analytical results obtained was quite large. Thus, the analytical results for DPhT obtained by ID–LC–ICP–MS were not adopted for the certification. The analytical results, with their uncertainties, obtained by both analytical techniques are summarized in Table 1. The values were calculated as mass fractions (based on dry mass). The combined standard uncertainties for the analytical results obtained by each method were calculated by use of Eqs. (1) or (2). The uncertainties related to the standard solutions (uncertainty of P and D in Eqs. (1) and (2)) were not combined into uc because the same reagents were used for both analyses. They were combined later when the uncertainties in the certified values were calculated.
Fig. 3GC–ICP–MS chromatograms of 118Sn and 120Sn obtained for the sample extract spiked with 118Sn-enriched butyltin and phenyltin compounds. The chromatogram of 118Sn was shifted 7 s and 2000 cps for clarityFig. 4GC–MS chromatograms of each measured mass obtained for the sample extract spiked with 118Sn-enriched butyltin and phenyltin compounds. The chromatogram for 118Sn was shifted 9 s and 500 cps for clarityFig. 5LC–ICP–MS chromatograms of 118Sn and 120Sn obtained for the sample extract spiked with 118Sn-enriched butyltin and phenyltin compounds. The chromatogram of 118Sn was shifted 9 s and 2500 cps for clarityTable 1Analytical results obtained by use of six combinations of extraction and measurement methods  Result±combined standard ua (μg kg−1 as Sn)ExtractionbMeasurementTBTDBTMBTTPhTDPhTUSEGC–ICP–MS44.3±1.151.2±0.965.9±1.17.8±1.43.4±0.3USEGC–MS43.8±1.750.7±0.866.6±1.75.8±1.53.9±0.4USELC–ICP–MS44.2±1.750.7±0.865.6±1.77.4±1.9–cMAEGC–ICP–MS43.8±1.552.1±0.968.2±1.27.0±1.02.9±0.13PFEGC–ICP–MS42.9±1.250.1±1.067.1±0.86.3±0.63.7±0.2MWEGC–ICP–MS47.0±1.352.5±1.267.6±0.88.7±1.74.4±0.8aCalculated by use of Eqs. (1) or (2)bUSE: ultrasonic extraction, MSE: mechanical shaking extraction, PFE: pressurized fluid extraction, MAE: microwave assisted extractioncTechnically invalid
The analytical results obtained by use of the six methods were in good agreement within the range of their uncertainties; this agreement may indicate there were no significant analytical biases between measurement and extraction techniques for all the analytes. Therefore, all the analytical results obtained were treated equally for calculation and evaluation of the certified values and their uncertainties.
Establishing certified values
The analytical results in Table 1 were combined to provide the certified values listed in Table 2. Certified values are the weighted means of these results from the six methods, where 1/ui (ui is the uncertainty of each result) was used as a weight. Certified values are available for the concentrations of MBT, DBT, TBT, DPhT, and TPhT, as tin.
Table 2Certified values and their uncertainties for mass fractions of organotin compounds in NMIJ CRM 7306-a TBTDBTMBTTPhTDPhTCertified value (mass fraction, μg kg−1 as Sn)4451676.93.4Relative standard uncertainty (%) Calibration solution ucal0.3%0.3%0.3%0.5%1.0% Analytical results uanal1.3%0.7%0.7%7.0%3.3% Between method umethod1.6%1.0%0.2%–– In-homogeneity sbb or ubb1.9%1.9%2.0%5.8%7.1%Combined uncertainty Relative (%)2.8%2.3%2.1%9.1%7.9% Absolute (μg kg−1 as Sn)1.31.21.40.60.3Expanded uncertainty U (k=2) (mass fraction, μg kg−1as Sn)3231.30.5
Uncertainty of the certified values
The uncertainties of the certified values included the combined effects of method imprecision, possible bias effects among methods, material inhomogeneity, and stability. Components of the uncertainty of each certified value are listed in Table 2. Because the same reagents for calibration solutions were used for all the measurements, uncertainty of the calibration solutions (ucal) was combined with the uncertainty of the certified values separately from the uncertainty of the analytical results, as is described above. The property value was the weighted mean of the two results, and so the combined uncertainty of each analytical result (uanal) was given by Eq. (5):
where xi are the results obtained by use of the six methods and wi are their weights. The between-method variance (umethod) was calculated from ANOVA on the data from the three techniques. The uncertainties derived from material inhomogeneity (ubb) were the estimates in the homogeneity study. As is described above, the butyltin and phenyltin compounds would be stable at lest five years at our storage temperature (−30 °C); we did not include the uncertainty from stability. The expanded uncertainty in each certified value is equal to U=kuc, where uc is the combined standard uncertainty, with coverage factor k=2, corresponding to 95% confidence intervals.
Comparison with other CRMs
The certified values for CRM7301-a and the CRM7306-a are summarized in Table 3. In comparison with the CRM7301-a, the uncertainties of the certified values for DBT and MBT in CRM7306-a are significantly smaller, despite the similar concentrations of the compounds. The main contributors to the uncertainty in the certified values for DBT and MBT in CRM7301-a were ubb and uanal [3]. The homogeneity of CRM 7306-a was improved by remixing after X-ray sterilization, and the reliability of the analytical methods used for certification was also better than those used for CRM7301-a. Thus, these improvements may lead to the small uncertainties.
Table 3Certified values for butyltin and phenyltin in MNIJ7306-a and other CRMs certified by use of SSIDMS (unit: µg kg-1 as Sn) TBTDBTMBTTPhTDPhTCRM7306-a44±351±267±36.9±1.23.4±0.5CRM7301-a44±456±656±6HIPA-178±9SOPH-1125±7174±9PACS-2890±1051047±64
Most recently, several sediment reference materials for butyltin analysis certified by use of SSID–MS have been issued by the National Research Council Canada (NRCC, Canada). The certified values of the CRMs are also summarized in Table 3. HIPA-1 and SOPH-1 were certified, and PACS-2 was recertified by NRCC. Their certified and re-certified values were based on results from the Comite Consultatif pour la Quantite de Matiere (CCQM) comparisons [23–25], in which we participated. These CRMs have certified values for DBT and TBT only, and their concentrations are higher than those in CRM7301-a and CRM7306-a.
The BCR 646 freshwater sediment for butyltins and phenyltins are also available from the European Commission Joint Research Centre (IRMM, Belgium), although SSID–MS was not used for the certification. The certified concentrations for BCR CRM646 are higher than those for CRM7306-a.
In Japan, typical butyltin and phenyltin concentrations observed in environmental monitoring are not as high as they used to be. Reported TBT concentrations in sediment were below 80 μg kg–1, as Sn, in environmental monitoring by the Ministry of the Environment Japan in 1998 [26]. By using these CRMs properly, a wide concentration range can be covered in routine laboratories.

certification
sediment
certified reference material
organotin compounds
species-specific isotope dilution


Chemotactic activity of extracellular nucleotideson human immune cells.

Purinergic P2 receptors are a class of plasma membrane receptors that are express in many tissues and are ligated by extracellular nucleotides [such as adenosine triphosphate (ATP), adenosine diphosphate (ADP), uridine 5–triphosphate (UTP) and uridine 5–diphosphate (UDP)], which are released as a consequence of cell damage, cell stress, bacterial infection or other noxious stimuli. According to the molecular structure, P2 receptors are divided into two subfamilies: P2X and P2Y receptors. The P2X receptors are ligand-gated channels, whereas P2Y receptors are G-protein-coupled seven-membrane-spanning receptors. Several studies indicate that nucleotides play an important role in immune response modulation through their action on multiple cell types, including monocytes, mast cells, dendritic cells, neutrophils, and eosinophils. Recent work by our group and others identified extracellular nucleotides as chemotaxins for various human immune cells, including eosinophils, neutrophils and dendritic cells. In this review, we summarise recent findings in this field and put forward a hypothesis on the role of P2 receptors in the early recruitment of human immune cells to the site of inflammation.
Introduction
Cell migration plays a key role in a wide variety of biological processes, such as embryogenesis, development, angiogenesis, haematopoiesis, immune response and inflammation. In inflammation and host defence, the targeted trafficking of immune cells to tissues and/or lymphoid organs is one of the essential steps. Migration of the different leukocytes is tightly controlled by chemokines. These chemotactic cytokines are secreted proteins with a molecular weight of 8–0 kDa, which direct cellular traffic along ingeniously regulated concentration gradient in the extracellular space. Based on amino acid alignments, chemokines are divided into four families. According to the position and the spacing of the first two conserved cysteines or the lack of them, these families are distinguished as either C, CC, CXC or CX3C. Until now, C and CX3C are composed of only one member each, lymphotactin and fractalkine, respectively, whereas CC and CXC each consist of numerous, well-characterised members [1, 2]. The chemotactic effects of these molecules are mediated due to their interactions with different specific serpentine receptors that span the plasma membrane seven times and belong to the G-protein-coupled receptor family.
Besides chemokines, many constitutive molecules can regulate the function of leukocytes and thereby modulate immune responses, e.g. following tissue damage, intracellular localised substances can be released into the extracellular space. For this reason, an increase of the extracellular concentration of certain molecules can be a very simple sign of cell damage. However, for constitutive molecules to function as chemotaxins, it is essential that they are recognised by immune system migration cells. In the past few years, evidence has accumulated strongly suggesting that nucleotides fulfil these requirements. They are present at high concentrations (5–0 mM) in the cytoplasm of all cells, whereas in the extracellular compartment, their concentration is in the nanomolar range. Nucleotides can be released into the extracellular space via nonlytic mechanisms through regulated transport; e.g. adenosine triphosphate (ATP) has been reported to be secreted by different cell types in a broad variety of conditions, such as shear stress, endotoxin stimulation, or at sites of platelet aggregation [3, 4] Hence, in tissues, nucleotides are able to generate concentration-dependent gradients, which can serve as chemotactic signals for different immune cells, causing migration.
P2 receptors
P2 receptors are subdivided on the basis of pharmacological, functional and cloning data into two families: the P2YR and P2XR [5–8]. P2YR are seven-membrane-spanning, G-protein-coupled receptors, and eight different P2YR subtypes have been cloned so far (P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13 and P2Y14) [5, 9, 10]. Activation of P2YR induces phospholipase C activation, inositol triphosphate generation, Ca2+ release from intracellular stores, and/or stimulation/inhibition of adenylate cyclase. Extensive pharmacological studies performed in P2Y transfected cells revealed that P2Y1, P2Y11, P2Y12 and P2Y13 selectively interact with ATP and/or adenosine diphosphate (ADP), whereas uridine 5–triphosphate (UTP) and uridine 5–diphosphate (UDP) are inactive [11–15]. In contrast the P2Y2, P2Y4 and P2Y6 subtypes are responsive to uridine nucleotides [16–18]. Whereas ATP and UTP activate P2Y2 with similar efficiency, UTP and UDP are most the potent agonists at P2Y4 and P2Y6, respectively [16–18]. In addition, it has been shown that P2Y14 specifically responds to UDP glucose and related sugar nucleotides but not to ATP, ADP, UTP or UDP [5, 9, 19, 20]. P2YR have been shown to modulate multiple cell function of various human immune cells, including cytokine release from dendritic cells, reactive oxygen metabolite production from neutrophils and eosinophils or chemokine release from airway epithelial cells [20–23].
P2XR are multimeric ligand-gated plasma-membrane ion channels activated by extracellular ATP and selective for monovalent and divalent cations [8, 24, 25]. At this time, seven different monomers have been cloned: the P2X1–P2X7 subtypes. Activation of P2XR leads to increased plasma membrane permeability to ions (Na+, K+, and Ca2+) and induction of apoptosis in human immune cells [22, 26] . In contrast to P2YR, the only currently known physiological ligand for all P2XR subtypes is ATP.
P2 receptors and migration of neutrophils
The first report that extracellular nucleotides can modulate human neutrophil function was a paper by Ward et al. showing that ATP and ADP can induce superoxide anion formation [27, 28]. Furthermore, it has been shown that human neutrophils or human promyelocytic HL60 cells respond to ATP, ATPγS and UTP with an increase in intracellular Ca2+ concentration via a pertussis toxin-sensitive G-protein receptor that coupled to the inositol phospholipid signaling system, suggesting involvement of P2Y subtypes [27, 29–33]. Activation of these intracellular signal-transduction systems is of great interest in the light of neutrophil recruitment to the site of inflammation, as increase in Ca2+ concentration is an important step in human neutrophil migration [34–36]. Furthermore, it has been shown that ATP also increases membrane expression of CD11b/CD18 and adhesion to albumin-coated polystyrene latex beads [37].Up-regulation of these molecules enhances the adhesion of neutrophils to other cells, e.g. between neutrophils and pulmonary endothelial cells, and could be of relevance for neutrophil migration across the vessel wall. However, after earlier observations suggesting that nucleotides might be chemoattractant for human neutrophils, it was not until the end of the 1990s that ATP and UTP were shown, by Verghese et al. [103] to induce actin polymerisation and chemotaxis in human neutrophils via the activation of P2Y2 (formerly known as P2U) receptor.
Surprisingly, the first reverse transcripaise polymerase chain reaction (RT-PCR) data revealed that human neutrophils express only P2Y4 and P2Y6 but not P2Y1 and P2Y2 receptors [38]. However, recent reports showed that human neutrophils also express messenger ribonucleic acid (mRNA) for the P2Y2 and P2Y11 receptor subtypes [39, 40]. Among the P2X receptors, so far, only the present of the P2X7 receptor has been shown by Northern blotting and immunocytochemistry [41]. Because increased extracellular nucleotide concentrations are associated with cell damage/injury and human neutrophils are the initial cell type found at tissue injury sites, the conclusion could be drawn that extracellular nucleotides among other mediators are involved in the recruitment of neutrophils to the site of inflammation.
P2 receptors and recruitment of human eosinophils
Besides neutrophils, eosinophils also express P2 receptors. Several studies showed that human eosinophils express mRNA for the following P2Y and P2X subtypes: P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y14, P2X1, P2X4 and P2X7 [23, 42–44]. Stimulation of P2 receptors expressed by eosinophils induces multiple cell responses, including production of reactive oxygen metabolites and secretion of eosinophil cationic protein (ECP) [23, 26].
Burgers and colleagues [45] showed that ATP, secreted by thrombin-activated platelets, was able to raise eosinophil intracellular Ca2+ concentration and made the cells chemotact towards platelets. These seminal studies were later confirmed by the identification of eosinophil P2Y and P2X receptors [42, 44] and the observations that nucleotides also induce up-regulation of adhesion molecule CD11b and actin polymerisation to important features involved in blood eosinophil recruitment to tissue [23, 42, 44].
In addition to direct chemotactic influences, ATP might also have indirect effects on eosinophil recruitment. We recently showed that ATP and UDP induce secretion of interleukin (IL)-8 by eosinophils [26]. This chemokine is a potent attractor for eosinophils themselves (and neutrophils), i.e. they are able to recruit more cells to inflammation sties. Increased secretion of IL-8 has been described in eosinophils from patients with bronchial asthma or atopic dermatitis [46]. Moreover, IL-8 concentration in bronchoalveolar fluids from asthmatic patients is increased significantly in comparison with that of healthy subjects [46]; therefore, one can suggest the involvement of different nucleotides in the direct or IL-8-mediated recruitment of eosinophils and thus in the development and maintenance of allergic diseases.
P2 receptors and mast cells
Mast cells are situated around blood vessels and nerves, especially at interfaces with the external environment, emphasising their role in immunity. They express several mRNAs that encode P2XR and P2YR subtypes [47, 48]. Human mast cells express P2X1 and P2X4, whereas the P2X7 receptor subtype is only expressed by human-cord-blood-derived mast cells when activated with anti-immunoglobulin (Ig)E [47]. Among the P2Y receptors, the presence of P2Y1, P2Y2, P2Y11, P2Y12 and P2Y13 subtypes were shown by RT-PCR [48]. Moreover, ATP and UTP enhance histamine release by human lung mast cells stimulated by cross-linkage of the fragment crystallisable (Fc)εRI [49]. This effect is attributed to the P2Y2 receptor. Data on chemotactic effects of nucleotides on mast cells are rare, but there is evidence that they might effect the migration of these cells. For example, McCloskey and colleagues showed that the nucleotides ADP, ATP and UTP are effective chemoattractants for rat-bone-marrow-cultured mast cells [50]. However, whether nucleotides can also induce migration of human mast cells remains to be elucidated
P2 receptors and lymphocytes
The first evidence for a role of extracellular nucleotides in human lymphocyte responses has been present for some time [51–53], but a systemic analysis of the expression and function of P2 receptors in human lymphocytes was only started at the end of the 1980s [30, 54–57]. Human B lymphocytes express both P2X and P2Y receptors [58, 59]. The presence of different P2Y receptors is indicated by the ability of ATP and many other nucleotides to trigger Ca2+ release from intracellular stores [60, 61] and the finding of P2Y1, P2Y2, P2Y4 and P2Y6 receptor-specific mRNA in lymphocytes [38]. Different studies suggest that human B cells express at least P2X7 receptor [22], but the identification of other P2X receptor subtypes is limited by the absence of specific antibodies [62, 63]. Nevertheless, confocal microscopy studies using anti-P2X polyclonal antibodies suggest the presence of P2X1, P2X2, P2X4 and P2X7 subtypes on human B lymphocytes [59]. However, despite the presence of functional P2 receptors, data about chemotactic effects on B lymphocytes induced by nucleotides are still missing.
Functional and pharmacologic studies revealed that human peripheral T lymphocytes express P2X-like ATP-activated channels, most likely P2X1, P2X4 and P2X7, [22, 64, 65]. Functional activity of the P2X receptors on T cells has been shown by a large influx of Na+ and Ca++ from the extracellular medium caused by ATP and 3–O-(4-benzoyl)benzoyl-ATP (Bz-ATP) [65].
The expression of P2Y receptors subtypes is still unclear, and whereas Baricordi and coworkers described a lack of functional P2Y receptors expression [65], recent studies using complete lymphocyte populations (T and B cells) could detected all the target genes for P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12 and P2Y13 [66]. However, a functional expression of different P2Y receptors on human T lymphocytes has not been proven.
Studies showing that activation of the P2X7 receptor by ATP and Bz-ATP induced shedding of CD23 and L-selectin from B and T lymphocytes of a B-chronic lymphocytic leukaemia (B-CLL) patient and from normal subjects, a classic effect of chemoattractants [67–70], leads to the assumption that the P2X7 receptor might be involved in the transendothelial migration of lymphocytes [71, 72]. But elegant experiments from Chen and coworkers using P2X7 antagonist in in vitro migration assays indicated that ATP is neither a chemoattractant that stimulates transmigration of lymphocytes nor an agonist that mediates the global L-selectin loss during transendothelial migration [73]. In any case a direct effect of extracellular nucleotides on T and B lymphocyte migration has not yet been investigated and, therefore, the role of nucleotides in the recruitment of lymphocytes remains unclear.
P2 receptor in monocyte/macrophages
Although the first report on a potential role of exogenous nucleotides on mouse macrophage function was a paper by Cohn and Parks from 1967 [74], a systemic investigation on human monocytes/macrophages was only started in the 1990s [30, 38, 75–78]. Freshly isolated blood monocytes have been shown to express mRNA for the following P2Y and P2X subtypes: P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13 P2X1, P2X4 and P2X7 [38, 66, 79]. However, other investigations found a lack of functional P2X7 in these cells, whereas the receptor appears during maturation of monocytes to macrophages [22]. Besides P2X7 receptors, human macrophages or macrophage cell lines have been described to express P2Y2, P2Y4 and P2Y6 receptors [30, 80, 81].
Activation of P2 receptors expressed by human monocytes/macrophages induces multiple cell responses, including increase of intracellular calcium concentration, induction of apoptosis, generation of reactive oxygen intermediates, NO generation and secretion of IL-1β, tumour necrosis factor (TNF)-α or IL-18 [22, 30, 77, 79, 82, 83]. In the light of migration, it is of great interest that extracellular ADP causes increased surface expression of MAC-1 (alpha M beta 2 integrin, CD11b/CD18) on monocytes [84] and that nucleotides can induce the adherence of monocytes to surfaces [38], which could be of relevance for monocyte migration across the vessel wall. Accordingly, migration of the mouse macrophage line J774 towards ADP has been demonstrated [50]. Interestingly, Warny et al. demonstrated that UDP activates IL-8 gene expression and IL-8 release in human monocytic cells [85]. Because IL-8 is a central mediator in inflammation and an important chemotactic factor for various cells, including neutrophils, eosinophils and CD16+ natural killer (NK) cells [46, 86–89], UDP [through its action on dendritic cells (DC)] might be indirectly involved in the recruitment of these cells to the side of inflammation. However, at this time, no information on the chemotactic activity of nucleotides on human monocytes/macrophages is available.
P2 receptors in dendritic cells
DCs are powerful antigen-presenting cells that circulate in the bloodstream or reside in peripheral tissues. They are characterised by a high antigen uptake capacity, recognition of constitutive or inducible endogenous ‘danger signals–provided by surrounding cells and a high responsiveness to chemotactic signals. The migratory ability of DCs is one of the main features in the initiation of immune responses. After acquiring antigens in the peripheral tissue, DCs migrate to the draining mediastinal lymph nodes to activate naive T cells [90–92]. Besides this ‘classic–action, some evidence also suggests that tissue-resident DCs are able to uptake tissue antigens and to migrate to the afferent lymph nodes, even in the absence of inflammatory conditions, thus contributing to tolerance maintenance [93, 94].
In the last few years, DCs came into the focus of researchers of the purinergic field, and the role of P2 receptors in DC migration came to the fore. RT-PCR analysis revealed that human DCs express a broad variety mRNAs for at least four subtypes of the P2X receptor family (P2X1, P2X4, P2X5, P2X7) and eight subtypes of the P2Y receptor family (P2Y1, P2Y2, P2Y4, P2Y6, P2Y11 P2Y12 P2Y13, P2Y14) [95–99].
First, studies by Liu et al. revealed that at least activation of the P2Y1, P2Y2 and P2Y4 mediates calcium release from intracellular storage. Furthermore, the observation that DCs redirect their dendrites towards a nearby patch pipette leaking ATP suggested that P2YR might mediate DC chemotactic response [96]. Indeed, ATP and UTP, probably via activation of the P2Y2, as well as ADP (via P2Y1?) turned out to be potent chemotactic stimuli for immature but not for mature DCs [100] In contrast, P2X receptor activation had only marginal chemotactic activity in both immature and mature DCs. Chemotaxis was paralleled by other intracellular signalling events, such as actin polymerisation and intracellular Ca2+ mobilisation. Recently, UDP could be added to the list of chemoattractant nucleotides, as it has been shown that UDP via binding to the P2Y6 receptor increased intracellular calcium, induced actin polymerisation and migration of immature, but again not mature, DCs [101]. The discrepancy between the responsiveness to extracellular nucleotides of immature and mature DCs could be explained by functional studies. They revealed a selective down-regulation of the Gi/o protein-coupled chemotactic P2Y receptor responsiveness during maturation. Surprisingly, immature and mature DCs expressed similar amounts of mRNA for the purinergic receptor subtypes P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2X1, P2X4 and P2X7 [100]. DC maturation encompasses a coordinated down-regulation of inflammatory chemokine receptors (CCR1, CCR2, CCR5 and CXCR1) and induction of CCR7 and CXCR4 [2]. In addition, during maturation, functional down-regulation of chemotaxis-regulating P2Y receptors are uncoupled to chemotaxis-associated signal transduction pathways. As a result, DCs lose sensitivity to inflammatory chemokines as well as to the nucleotides ATP, ADP, UTP and UDP.
In addition, nucleotides can modulate the migration of DCs to chemokines, as immature and matured DCs stimulated with ATP gain the ability to migrate in response to CXC ligand (L)12 and CCL12 [102]. However, in contrast, Schnurr et al. reported that ATP through P2Y11 signalling could inhibit CCL21, induce migration of immature and mature monocyte-derived DCs and CD1a+ dermal DCs but not of CD1c+ peripheral blood DCs or IL-3R+ plasmacytoid DCs [98]. This controversy could be due to differences in the blood donors or preparation of monocyte-derived DCs in the different laboratories.
Besides direct chemotactic effects on DCs, extracellular nucleotides are also involved indirectly via their action on DCs in the trafficking of other leukocytes through the release of chemokines. For example, ATP up-regulates the constitutive production of CCL22 [macrophage-derived chemokine (MDC)] and inhibits the lipopolysaccharide (LPS)-induced secretion of CXCL10 (IP-10) and CCL5 (RANTES), resulting in selectively impaired recruitment of type 1 but not type 2 T cells, suggesting a nucleotide-mediated communication between DCs and T cells an important event during antigen presentation in vivo [102]. In accordance, UDP can enhance the LPS-mediated release of chemotactic factor IL-8 via P2Y6 from mature DCs [101], so again, UDP might be indirectly involved in the recruitment of neutrophils, eosinophils and CD16+ NK cells to the side of inflammation.
In sum, activation of DCs by extracellular nucleotides leads to multiple cell responses, which results in a direct migration of DCs and also maybe indirect (DC-mediated) recruitment of other immune cells to the site of inflammation.
Conclusion
Over the last few years, several studies have implied that extracellular nucleotides which are actively released or diffuse out of mechanically stressed, infected or injured cells might be involved in the early recruitment of immune cells to the site of inflammation/cell damage. In these in vitro experiments, it has been shown that extracellular nucleotides (mainly by activating P2YR subtypes) are direct chemoattractants for human neutrophils, eosinophils and DCs and/or they can modulate the chemokine production of eosinophils, monocytes and DCs, which might then influence the migration capacity of other immune cells. However, whether nucleotides play a role in the migration of immune cells in vivo still remains to be elucidated.

nucleotides
immune cells
chemotaxins
purinergic receptors