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Organotin compounds: their analyses and effect on model biomembranes Nwata, Basil Ugwunna 1994

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ORGANOTIN COMPOUNDS:- THEIR ANALYSES AND EFFECT ONMODEL BIOMEMBRANESBYBASIL UGWUNNA NWATAB.Sc (Hons), University of Ilorin, Nigeria, 1981M.Sc, University of Ibadan, Nigeria 1984M.Sc, University of British Columbia, Canada 1989A THESIS SUBMITrID IN PARTIAL FULFILLMENT OF THEREQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHYIN THE FACULTY OF GRADUATE STUDIESDEPARTMENT OF CHEMISTRYWe accept this thesis as conforming to the required standardCOLUMBIAJUNE 1994© Basil Ugwunna Nwata, 1994In presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)__________________Department of C/17’1’ i7’The University of British ColumbiaVancouver, CanadaDate ditW4 i’?DE-6 (2)88)11ABSTRACTStudies involving the analyses of organotin compounds in marine organismsof British Columbia, and the effect of organotin compounds on the permeability ofmodel biological membranes are presented in this thesis.Analysis of organotin compounds by gas chromatography-selected ionmonitoring mass spectrometry (GC-MS SIM) affords a very specific technique for theidentification and quantitation of organotin compounds, by using the peculiar isotopepattern for tin compounds. This methodology is therefore able to distinguishorganotin compounds from other compounds that may co-elute with them from thegas chromatograph.Although some British Columbian locations such as Hastings Arm, Alice Arm,etc showed no organotin contamination, the major organotin pollutants found forsome coastal areas such as Denman Island, Dundas Island, etc were tributyltin anddibutyltin species. The butyltin body content for Blue mussels in the contaminatedareas range from 14.4 to 37.3 ng/g (wet wt as Sn) for tributyltin and 6.7 to 67.3 ng/g(wet wt as Sn) for dibutyltin species. Dicyclohexyltin levels of 3.5 ng/g and 21.3 ng/g(wet wt as Sn) were found only at Wreck Beach Vancouver, and Anyox respectively.The effect of organotin compounds on egg phosphatidyicholine (EPC)liposomes or organotin-EPC liposomes, were established by studing the efflux of aprobe compound; dimethylarsinic acid (DMA) trapped inside these liposomes, byusing 1H NMR spectroscopy. The probe compound at pH 7.4 exists as two chemicalspecies; DMAH and DMK which are capable of diffusing from these liposomes.111When the organotin compounds were added externally to the EPC liposomes,tributyltin chloride caused an increased permeability of the liposomes, which waslinearly dependent on the concentration of the externally added tributyltin chloridesolution. Monobutyltin trichioride decreased the permeability coefficient of DMAHto the EPC liposomes from 1.7 x 10-8 to 4 x i0 cm/s, while trimethyltin cationfacilitated the efflux of DMK from the liposomes.For TBT-EPC liposomes formed by a mixture of tributyltin chloride and EPC,the efflux of DMA from these liposomes was facilitated by the tributyltin cation onlyif the liposomes were not in contact with externally added tributyltin chloridesolution. When in contact with externally added tributyltin chloride, the ability of thetributyltin cation to act as carrier for DMK was lost. The activation energy for thepassive efflux of DMAH from TBT-EPC liposomes varied from 52.3 to 64.4 kJ/moldepending on the tributyltin content of the liposome.For the monobutyltin trichloride-EPC liposomes (MBT-EPC), themonobutyltin cation did not exhibit any ability to act as carrier for DMA irrespectiveof whether it was externally added to the liposomes or not. The DMAH speciespermeate by passive diffusion with activation energy of 106.8 to 121.5 kJ/mol.A modified batch hydride generation-graphite furnace atomic absorptionspectrophotometric method (HG-GFAAS) is described for total tin determination.In this method, tin hydride was adsorbed and pre-concentrated on graphite furnacetubes pre-coated with palladium or sodium tungstate matrix modifiers, prior to theiratomization in the graphite furnace.ivTABLE OF CONTENTSAbstractTable of contents ivList of Tables xivList of Figures xixList of Abbreviations xxiiiAcknowledgements xxviDedication xxviiChapter 1General Introduction 11.1- Historical background of the organotin compounds 11.2 Industrial applications and use of organotin compounds 11.3 Need for a chemical antifouling agent 31.3.1 Contact leaching antifouling paints 51.3.2 Ablative formulation 51.3.3 Self polishing co-polymer paints 61.4 Toxicity of organotin compounds 61.5 Metabolism and behaviour of tributyltin compoundsin the environment 91.6 Analytical methods for organotin compounds 111.6.1 Molecular spectrophotometry and spectrofluorimetric techniques 121.6.2 Electrochemical techniques 13V1.6.3 Atomic spectrometry 141.6.4 Gas chromatography 171.6.4.1 Hydride generation gas-chromatography (HG-GC) 171.6.4.2 Conversion to tetraalkyltin compounds 191.6.4.2(a) Gas chromatography with flame photometricdetection (GC-FPD) 201.6.4.2(b) Gas chromatography with atomic absorptionspectrometric detection (GC-AAS) 211.6.4.2(c) Gas chromatography with mass spectrometricdetection (GC-MS) 211.6.5 Liquid chromatography (LC) 211.6.6 Thin layer chromatography (TLC) and high performancethin layer chromatography 241.7 Tributyltin and government regulations 251.8 Objectives and scope of the present study 26Chapter 2 Speciation and quantitation of butyltin and cyclohexyltincompounds in marine organisms by usingcapillary column GC-MS SIM 292.1 Introduction 292.2 Experimental 302.2.1 Instrumentation 302.2.1.1 Gas chromatography (GC) 30vi2.2.1.2 NMR and mass spectrometry 302.2.1.3 Gas chromatography-mass spectrometry (GC-MS) 312.2.1.4 Mechanical shaker and blender 312.2.2 Materials and reagents 322.2.3 Synthesis of standard organotin compounds 332.2.3.1 Synthesis of tributylmethyltin 332.2.3.2 Synthesis of dibutyldimethyltin 332.2.3.3 Synthesis of tricyclohexylmethyltin 342.2.3.4 Synthesis of dicyclohexyldimethyltin 342.2.3.5 Synthesis of the internal standards 342.2.3.5(a) Synthesis of tetrapropyltin 342.2.3.5(b) Direct synthesis of deuterated internal standards 352.2.3.6 Synthesis of methylmagnesium iodide 362.3 Analytical procedure 362.3.1 Gas chromatography 362.3.1.1 Establishment of elution profile and retention data 362.3.1.2 Suitability of tetrapropyltin as internal standard 372.3.1.3 Suitability of(C42H9)3SnCH and (C42H9)Sn(CH3 asinternal standards 372.3.2 Low resolution mass spectrometry 382.3.3 GC-MS retention data, calibration curves and precision 382.3.4 Recovery studies 39vi’2.3.5 Extraction of the organotin compounds from marine animals 412.4 Results and discussion 432.4.1 Characterization of the standard tetraorganotin compounds 432.4.2 Fragment ions and intensities of the standard tetraorganotincompounds 482.4.3 GC-MS elution profile and masses of selectedfragment ions used for selected ion monitoring 512.4.4 Suitability of tetrapropyltin as internal standard asstudied by gas chromatography 532.4.5 Detection limit, calibration curves, andprecision for the GC-MS SIM analysis 572.4.5.1 Detection limit and calibration curves obtainedby GC-MS SIM 572.4.5.2 Precision of the GC-MS SIM method 622.4.6 Recovery studies on the extraction procedure 632.4.7 Organotin concentrations in some marine organismsof British Columbia, Canada 672.4.7.1 Organotin concentrations in oysters 672.4.8 Spread of organotin compounds in the Canadian environment 712.4.9 Organotin concentrations in various organisms from thesame locations 79vii’2.4.10 Distribution of organotin compounds in marine animalsstudied over a period of three years 81Chapter 3 Effect of tributyltin chloride, monobutyltin trichioride andtrimethyltin hydroxide on the permeability of eggphosphatidyicholine liposomes 833.2 Dimethylarsinic acid (DMA) as a probe for studying the effectof organotin compounds on the membranes of liposomes 843.3 Liposomes as models for biological membranes 853.4 Types of liposomes and methods of preparation 873.4.1 Multilamellar vesicles (MLVs) 883.4.2 Small unilamellar vesicles (SUVs) 883.4.3 Large unilamellar vesicles (LUV5) 893.5 Transport processes in membranes 903.5.1 Simple or passive diffusion 903.5.2 Facilitated diffusion 913.5.2.1 Solute translocation through channels 923.5.2.2 Translocation through carriers 923.5.3 Active transport 933.6 Solute transport across liposomal membranes 943.6.1 Transport of non-ionic solutes 943.6.2 Transport of ions 97ix3.7 Properties of liposomes capable of yielding investigativeinformation 993.8 Butyltin compounds: the need for the present study 993.9 Theoretical description of the diffusion experimentapplicable to NMR spectrometry 1013.9.1 Passive diffusion 1013.9.2 Facilitated diffusion 1013.10 Experimental 1113.10.1 Instrumentation 1113.10.1.1 Nuclear magnetic resonance spectrometry (NMR) 1113.10.1.2 Lipid extruder and membrane filters 1113.10.1.3 UV-Visible spectrophotometry 1113.10.2 Chemicals and reagents 1123.10.3 Preparation of large unilamellar vesicles (LUVs)from egg phosphatidyicholine (EPC) and theencapsulation of dimethylarsinic acid 1123.10.4 Preparation of butyltin-EPC LUVs and theencapsulation of DMA 1143.10.5 The NMR water suppression and spectral acquisitionconditions for DMA efflux from EPC andbutyltin-EPC liposomes 116x3.10.6 Determination of phospholipid concentrationsby phosphorus assay 1173.10.6.1 Extraction of phospholipid from liposomes prior tophosphorus determination 1173.10.6.2 Lipid concentration determination 1173.10.7 Processing of the NMR spectra 1183.10.8 Analysis and treatment of data 1203.10.8.1 Determination of rate constants and mode of permeation 1203.10.8.2 Determination of permeability coefficients 1243.11 Results and Discussion 1253.11.1 The use of DMA as a probe in permeability studies of EPCliposomes in the presence and absence of organotincompounds in the extraliposomal aqueous compartment 1253.11.2 Effect of organotin concentration on the efflux of DMA 1353.11.3 Efflux of DMA from tributyltin chloride-EPC liposomes(with tributyltin chloride absent in the extraliposomalcompartment) 1393.11.4 Efflux of DMA from monobutyltin trichloride-EPC liposomes 1453.11.5 Effect of the butyltin chloride concentrations of theliposome on permeability properties of TBT-EPC andMBT-EPC liposomes 149xi3.11.6 Effect of temperature on the permeability oforganotin-EPC liposomes 1513.11.7 Activation energies for the permeation ofbutyltin chloride-EPC liposomes 1533.11.8 Relevance of this NMR study to the enviromnental toxicity of.butyltin compounds 158Chapter 4 Hydride generation methods of atomic absorptionspectrophotometry for total tin determination 1604.1 Introduction 1604.2 Experimental 1624.2.1 Instrumentation 1624.2.1.1 Continuous hydride generation atomicabsorption spectrophotometry (HG-AAS) 1624.2.1.2 Batch hydride generation-graphite furnaceatomic absorption spectrophotometry (HG-GFAAS) 1644.2.2 Materials and Reagents 1654.2.3 Methodology for HG-AAS 1664.2.3.1 Continuous hydride generation method (HG-AAS) 1664.2.3.2 Batch hydride generation-graphitefurnace method (HG-GFAAS) 1664.2.4 Preparation of matrix modifiers and standard tin solutions 1684.2.4.1 Preparation of palladium modifier 168XII4.2.4.2 Preparation of sodium tungstate modifier 1654.2.4.3 Preparation of standard tin solutions 1694.2.5 Optimum concentration of reagents used in HG-AAS 1694.2.6 Use of L-cysteine to remove interferences 1704.2.6.1 Optimum concentration of L-cysteine required toremove interferences 1714;2.7 Optimum conditions for the batch HG-GFAAS 1714.2.7.1 Optimization of reagent concentrations for HG-GFAAS 1724.2.7.2 Optimization of the trapping temperatures and trappingtime for tin hydride in the graphite furnace tube 1724.2.8 Treated graphite furnace tubes:- coating the graphitefurnace tubes with solutions of sodium tungstateand palladium modifiers 1724.2.8.1 Optimum modifier treatment of graphite furnace tubes 1734.2.8.2 Calibration curves for the HG-GFAAS method 1734.3 Sample digestion and preparation 1744.4 Results and Discussion 1754.4.1 Optimum concentrations of sodium borohydride and HC1necessary for the production of stannane in thecontinuous hydride generator 1754.4.2 Optimum concentration of L—cysteine required to eliminateinteferences in HG-AAS 177XIII4.4.3 HG-AAS determination of total tin in oysters andstandard reference material (Tort 1) 1794.4.4 Batch hydride generation-graphite furnaceatomic absorption spectrophotometry (HG-GFAAS) 1824.4.4.1 Optimum concentrations of reagents needed for tinhydride production in the HG-GFAAS method 1844.4.4.2 Optimum flow rate of sodium borohydride intothe batch hydride generator 1854.4.4.3 Optimum temperature for trapping tin hydridein the pre-treated graphite furnace tubes 1864.4.4.4 Optimum trapping time 1874.4.4.5 Pre-treatment of graphite furnace tubes with modifiers 1884.4.4.6 Determination of total tin content of a standardreference material by the HG-GFAAS method 192Chapter 5 Summary and conclusions 195References 202Appendix A Map of locations sampled for organotin pollution 220Appendix B The NMR spectral acquisition and water suppression parametersfor the efflux of dimethylarsinic acid from liposomes 221Appendix C Michael is-Mentons equations for enzyme kinetics 223Appendix D Wet ashing apparatus with air cooled reflux condenserused for digestion of marine animals 224xivLIST OF TABLES PAGE2.1 119Sn NMR chemical shifts for the standardorganotin compounds 442.2 Major fragment ions of tributylmethyltin 482.3 Major fragment ions for dibutyldimethyltinand tricyclohexylmethyltin 492.4 Major fragment ions for dicyclohexyldimethyltinand tetrapropyltin 502.5 Retention time and retention time window used forGC-MS SIM analysis 512.6 Fragment ions and masses used to detect and quantitateeach organotin compound in GC-MS SIM 532.7 Regression data for graphs obtained with variousconcentrations of the internal standard tetrapropyltin 542.8 Detection limits for organotin compounds by GC-MS SIM 602.9 Calibration equations used for the quantitation ofenvironmental samples by GC-MS SIM 612.10 Precision of the GC-MS SIM 632.11 Recovery of organotin compounds spiked into Shrimp 642.12 Organotin concentrations in the oyster Crassostrea gigasfrom some coastal areas of British Columbia 682.13 Organotin concentration, spread and speciation in somexvlocations of British Columbia 722.14 Some organotin concentrations reported for theBlue mussels Mytilus edulis 772.15 Organotin concentrations in the Blue musselMytilus edulis converted to tg/g wet wt asorganotin cation 782.16 Organotin distribution in marine animals from Camano SoundBritish Columbia 792.17 Organotin distribution in marine animals fromTasu Sound British Columbia 802.18 Organotin body burden for soft shell clams fromQuatsino Sound, British Columbia studied overa period of three years 813.1 Efflux data for the diffusion of DMAH from EPCliposomes in the absence of organotin compounds 1263.2 Effect of 33.2 .LM tributyltin chloride on theefflux of DMAH from EPC liposomes 1273.3 Effect of 33.2 iM monobutyltin trichloride on theefflux of DMAH from EPC liposomes 1313.4 Data for the efflux of DMA from EPC liposomes inthe presence of 33.2 M trimethyltin hydroxide 1353.5 Effect of tributyltin chloride concentration onxviDMAH efflux 1363.6 Effect of monobutyltin trichloride concentration on theefflux of DMAH 1383.7 Diffusion parameters for the efflux of DMA fromtributyltin chloride-EPC liposomes by a mixture of passiveand facilitated diffusion 1413.9 Parameters for the efflux of DMAH from TBT-EPC Bliposomes in the presence of externally added tributyltinchloride (16 iiM) 1453.9 Permeability data for efflux of DMAH from MBT-EPC Bliposomes (with monobutyltin trichioride absent in theextraliposomal compartment) 1483.10 Permeability data for efflux of DMAH from MBT-EPC Bliposomes (with monobutyltin trichioride present in theextraliposomal compartment) 1483.11 Effect of tributyltin chloride concentration on thepermeability of tributyltin chloride-EPC liposomes withtributyltin chloride also present in the extraliposomalvolume 1503.12 Effect of monobutyltin trichloride concentration on thepermeability of monobutyltin trichloride-EPC liposomes withmonobutyltin trichioride also present in the extraliposomalxviivolume 1503.13 Effect of temperature on the permeability properties ofTBT-EPC A liposomes 1513.14 Effect of temperature on the permeability properties ofTBT-EPC B liposomes 1513.15 Effect of temperature on the permeability properties ofMBT-EPC A liposomes 1523.16 Effect of temperature on the permeability properties ofMBT-EPC B liposomes 1523.17 Effect of tributyltin chloride content of liposome onthe activation energy for efflux of DMAHfrom TBT-EPC liposomes 1553.18 Effect of monobutyltin trichioride content of liposomeson the activation energy for efflux of DMAHfrom MBT-EPC liposomes 1553.19 Arrhneius pre-exponential factor for DMAH effluxfrom TBT-EPC liposomes 1573.20 Arrhneius pre-exponential factor for DMAH effluxfrom MBT-EPC liposomes 1574.1 Graphite furnace atomization program for tindetermination by HG-GFAAS 1674.2 Operating conditions for the continuous hydride generationxviii-atomic absorption spectrophotometry (HG-AAS) 1704.3 Total tin content of samples analyzed by theHG-AAS method 1814.4 Reagent ratios needed to maximize tin hydride generation 1844.5 Comparison of palladium and sodium tungstate-treatedgraphite furnace tubes showing the atomic absorbanceof tin hydride generated from 14 g/mL tin solution 1924.6 Total tin content of a standard reference materialTort 1 obtained by different authors 1934.7 Comparison of figures of merit obtained with the twoatomic absorption spectrophotometric methods used inthis study 194xixLIST OF FIGURES PAGEFigure 2.1 Flow diagram for the extraction of organotin from marineanimals 42Figure 2.2 Mass spectra (El) of tricyclohexylmethyltin 45Figure 2.3 1H NMR spectra of dicyclohexyldimethyltin 46Figure 2.4 Mass spectra (El) of dicyclohexyldimethyltin 47Figure 2.5 GC-MS elution profile of the standard tetraorganotincompounds 52Figure 2.6 Effect of internal standard concentration on the linearity ofcalibration curves for tributylmethyltin 55Figure 2.7 Effect of internal standard concentration on the linearityof calibration curves for tributylmethyltin 56Figure 2.8 GC-MS calibration curves for (a) dibutyldimethyltinand (b) tributylmethyltin 58Figure 2.9 GC-MS calibration curves for (a) dicyclohexyldimethyltinand (b) tricyclohexylmethyltin 59Figure 2.10 Selected ion current chromatogram of standard organotincompounds spiked into shrimp 65Figure 2.11 Mass spectra of peak A in Figure 2.10 65Figure 2.12 Mass spectra of (i) peaks C and (ii) peak D in Figure 2.10 66Figure 2.13 (a) Selected ion current chromatogram of extract from Bluemussel from Wreck Beach, Vancouver. (b) Mass spectra ofxxpeak D in Figure 2.13(a) 76Figure 3.1 Structure of a phospholipid (phosphatidyicholine) 86Figure 3.2 Liposome 87Figure 3.3 Schematic diagram of facilitated diffusion (efflux)mediated by a carrier 93Figure 3.4 Passive diffusion of a permeant HA across a liposomalmembrane 96Figure 3.5 Proposed mechanism of tributyltin mediated efflux ofdimethylarsinate (DMK) from a liposome and the equilibriaof the carrier-permeant interactions 103Figure 3.6 1H NMR spectra of DMA as it diffuses out of EPC liposomes 119Figure 3.7 Log plot for the efflux of DMA from EPC liposomes 121Figure 3.8 Chemical species of dimethylarsinic acid present at pH 7.4 123Figure 3.9 Efflux of DMA from EPC liposomes (organotin compoundsare absent in the extraliposomal comparment) 126Figure 3.10 Time course for the efflux of DMA from EPC liposomes(tributyltin chloride present in the extraliposomal compartment) 128Figure 3.11 Time course for the efflux of DMA from EPC liposomes(monobutyltin trichloride present in theextraliposomal compartment) 128Figure 3.12 Log plot of DMA efflux from EPC liposomes (33.2 Mtrimethyltin hydroxide present in extraliposomal volume) 133xxiFigure 3.13 Time course for DMA efflux from EPC liposomes(33.2 jM trimethyltin hydroxide present in extraliposomalvolume) 133Figure 3.14 Effect of tributyltin chloride concentration on thepermeability of EPC liposomes (tributyltin chloride wasadded into the extraliposomal compartment) 137Figure 3.15 Effect of monobutyltin trichioride on the permeabilityof EPC liposomes (monobutyltin trichioride was added intothe extraliposomal compartment) 137Figure 3.16 Log plot of DMA efflux from TBT-EPC C liposomes 140Figure 3.17 Time course for DMA efflux from TBT-EPC C liposomes 140Figure 3.18 Contribution of passive and facilitated diffusion to theefflux of DMA from TBT-EPC liposomes of differenttributyltin chloride composition 142Figure 3.19 Log plot for efflux of DMA from TBT-EPC C liposomes when16.7 iM tributyltin chloride is present in theextraliposomal compartment 143Figure 3.20 Time course for efflux of DMA from TBT-EPC C liposomeswhen 16.7 iM tributyltin chloride is present in theextraliposomal compartment 144Figure 3.21 Log plot of DMA efflux from MBT-EPC B liposomes 147Figure 3.22 Time course for DMA efflux from MBT-EPC B liposomes 147xxiiFigure 3.23 Arrhenius plot for DMAH efflux from TBT-EPC B liposomes 154Figure 3.24 Arrhenius plot for DMAH efflux from MBT-EPC B liposomes 154Figure 4.1 Schematic diagram of the apparatus used for the HG-AASmethod 163Figure 4.2 Schematic diagram of the hydride generator used forthe HG-GFAAS method 163Figure 4.3 Effect of sodium borohydride and HC1 concentrations onthe absorbance of tin hydride produced from 4 JLg/mL tinsolution 176Figure 4.4 Effect of the concentration of L-cysteine on the absorbanceof tin hydride 178Figure 4.5 Effect of sodium borohydride flow rate on absorbance 185Figure 4.6 Effect of trapping temperature on the atomic absorbanceof tin hydride 187Figure 4.7 Effect of trapping time on absorbance 188Figure 4.8 Effect of sodium tungstate concentration on absorbance 191Figure 4.9 Effect of palladium on absorbance 191xxiiiLIST OF ABBREVIATIONSb.p Boiling pointCalcd Calculated valuecm CentimeterConc ConcentrationContd ContinuedDMA Dimethylarsinic acid or equilibrium mixture ofDMA and DMAHDMK Dimethylarsinate; negatively charged species ofdimethylarsinic acid present in solution at pH 7.4DMAH Undissociated dimethylarsinic acid present in solution atpH 7.4El Electron ionizationEPC Egg phosphatidyicholineGC Gas chromatograph/chromatographyGC-MS Gas chromatography with mass spectrometric detectionorGas chromatograph coupled to a mass spectrometerGC-MS SIM Gas chromatography with mass spectrometric detectionin the selected ion monitoring mode.GFAAS Graphite furnace atomic absorption spectrophotometryh hourxxivHEPES N-2-Hydroxyethylpiperazine-N’-2-ethanesulphonic acidHG-AAS Hydride generation atomic absorption spectrophotometryHG-GFAAS Hydride generation-graphite furnace atomic absorptionspectrophotometryHz Hertzi.d internal diameterLUVs Large unilamellar vesiclesM Molar (mol L1)MBT Monóbutyltin trichiorideMBT-EPC liposome Liposome formed by a mixture of monobutyltintrichioride and egg phosphatidyicholinemm minutesMS Mass spectrometer/spectrometryND Not detectedNMR - Nuclear magnetic resonance spectrometer/spectrometryppb Parts per billionppm Parts per millionpsi Pounds per square inchRSD Relative standard deviationSIM Selected ion monitoringTBT Tributyltin chlorideTBT-EPC liposome Liposome formed by a mixture of tributyltin chloride andxxvegg phosphatidyicholineiris Tris(hydroxymethyl)aminomethaneTSP Deuterated 3-(trimethylsilyl) propionic acid sodium saltv/v Volume to volume ratiow/v Weight to volume ratioxxviACKNOWLEDGEMENTS.I wish to express my gratitude to my research supervisor Professor W.R.Cullen for his guidance and interest in this research, and for my financial support.I would like to thank Dr Gunther Eigendorf for his advice on massspectrometry, and for his kindness towards me.I am also grateful to Madiba Saidy, Ryan Males, and the following people:Dr. ‘s Christopher Harrington, Bruce Todd, Roshan Cader, Kian Pang for their helpin the various stages of this thesis, and to the past and present members of ProfessorCullen’s research group for many helpful discussions.My gratitude also goes to the following people; Ms Lina Madilao, Mr SteveRak, Kim Wong and Gary Hewitt for their help in the various technical aspects ofthis work, and to Dr P.R. Cullis and Professor F.G. Herring for the use of theirfacilities.I am thankful to the Department of Chemistry, University of British Columbia,Canada for financial support and provision of research facilities.I also wish to express my sincere gratitude to my mother, brothers and sisters,and brother-Inlaw Dr A.N. Ewunonu, for their encouragement to me.To my wife Joyce, and children; Edoziem and Chima, I owe you muchgratitude for the encouragement and moral support you gave me.xxviiDEDICATIONThis thesis is dedicated to the memory of my beloved fatherChief Nelson Ukachi Nwata.Your departure before the task could be completed was very painful, but the dailyremembrance of you is my source of inspiration. Rest in perfect peace.1CHAPTER 1GENERAL INTRODUCTION.1.1. HISTORICAL BACKGROUND OF THE ORGANOT1N COMPOUNDS.The synthesis of diethykin diiodide by Frankland’ in 1849, marked theintroduction of a new class of compounds that were later to occupy a significantposition in industry and agriculture. By definition, organotin compounds are thosecompounds that have a carbon-tin covalent bond in the molecule. Progress in thechemistry of organotin compounds was enhanced by the discovery of Grignardreagents which made possible the production of a variety of organotin compoundsof formula R4Sn, from which lower alkyl- or aryl- tin compounds could easily bemade.The first industrial application of organotin compounds was made in 1936,when Yngve of the Carbide and Carbon Chemical Company, U.S.A., discovered theheat stabilizing effect of organotin compounds on polyvinyl chloride (PVC) and otherchlorinated hydrocarbon polymers2.The organotin compounds that have been founduseful in this application are the mono and dibutyltin compounds and the dioctyltincompounds.1.2 INDUSTRIAL APPLICATIONS AND USE OF ORGANOTENCOMPOUNDS.The organotin compounds find a wide range of use in the manufacturing2industry, agriculture and medicine. Because of their very low toxicity, the dioctyltinderivatives are used as stabilizers for food packaging polymers. Organotin compoundsare also used for cold curing of silicone rubber and as polymerisation catalysts; forexample, butyichiorotin dihydroxide3.Dibutyltin diacetate is a catalyst for flexiblefoams. Some organotin dihalides having the formula R2SnXL (R=ethyl or phenyl,X=chloride or bromide, L2= o-phenanthroline or 2-(2-pyridyl) benzimidazole exhibitanti-tumOur and anti-herpes activity in vitro4.Dibutyltin dilaurate is also effective inthe removal of intestinal worms in poultry3.Dicyclohexyltin derivatives of dipeptideshaving the formula Cy2SnL (L=glycylglycine, glycylalanine, glycyiphenylalanine andglycyltyrosine) exhibit high cytotoxicity in vitro to breast cancer cells5.The triorganotin compounds are the most important of the organotincompounds in agricultural applications. For example, tricyclohexyltin compounds areeffective as miticides and and possess marked acaricidal action against plant-feedingmites, but have very little effect on predacious mites and insects3. They also havebeen reported as antifeedants for the Gypsy moth Porthetria dispar6. The pesticidePlictran® marketed by Dow Chemical Company has tricyclohexyltin hydroxide as theactive ingredient7,and has been used in Canada for crop protection on apples andpears8. Peropal®, a pesticide marketed by Bayer AG has 1-tricyclohexyltin-1,2,4-triazole as the active ingredient.The triphenyltin compounds show antifungal activity. The fungicide Brestan®marketed by Hoechst A.G, Germany, contains triphenyltin acetate and has been usedagainst a broad range of fungal organisms in sugar beet and potatoes3. Du-Ter®, a3fungicide marketed by Philips-Duphar, Holland contains triphenyltin hydroxide asthe active ingredient3. The use of triphenyltin hydroxide as an antifeedant for theColorado beetle Leptinotarsa decemlineata has also been reported9.The tributyltin compounds are fungicides, algaecides and slimicides, and havebeen widely used as wood preservatives. Presently, their major use is in marineantifouling paint formulations for protecting the hulls of ships and boats from algae,fungi, sponges, molluscs, barnacles, diatoms, shipworms etc. Such fouling has theeffect of increasing weight and drag, causing the ship to consume more fuel tomaintain speed. The tributyltin compounds used in marine paint formulations arebis(tributyltin) oxide, bis(tributyltin) dodecenylsuccinate, bis(tributyltin) suiphide,tributyltin fluoride, tributyltin resinate, tributyltin methacrylate, bis(tributyltin)adipate. Unfortunately the tributyltin compounds are the major organotin compoundsof concern from the point of view of environmental marine pollution. When usedas antifouling agents, they pollute the marine environment. They do not remainlocalized but spread throughout the marine environment causing considerableproblems such as imposex in marine gastropods’° and the deformation and declineof oyster stock11.1.3 NEED FOR A CHEMICAL ANTIFOULING AGENT.The nuisance caused by the growth of unwanted marine organisms is a majorproblem in the maritime industry. One of the early approaches taken to preventfouling in wooden ships, was the use of copper metal sheathing12 on the ship’s hulls.4This achieved moderate success in the control of fouling. In steel ships, the use ofcopper metal sheathing is not appropriate because of the severe galvanic corrosionof steel when in contact with copper and sea water12. The usual method of foulingprevention in steel ships is to use chemical agents which are usually incorporated inthe paints used to paint the hulls. These antifouling paints act by releasing biocides,which kill the larvae and spores of any marine animals and plants attempting to settleon the ship’s hull.Among the early biocides employed for this purpose was cuprous oxide12.Cuprous oxide exhibits a wide spectrum of toxicity to animals, but many plants areresistant to it. Continuous use of the copper oxide results in the formation ofinsoluble greenish salts within the surface layers of the paint film. The build up ofthese salts on the surface prevents further controlled release of fresh biocide. Thisprocess limits the life time, and the efficiency of the paint. The search for biocidesto boost the performance of cuprous oxide led to the screening of the organotincompounds for biocidal activity. Tributyltin compounds were found to be suitablebiocides because they exhibit low mammalian toxicity, but high toxicity to fungi andalgae at low concentrations. They are also typically colorless and can therefore beincorporated into brightly colored paints.In the course of searching for efficient ways to deliver tributyltin compoundsto the target organisms, the following antifouling paint formulations have becomeavailable.51.3.1 Contact leaching antifouling paints.In this design, the antifouling system is composed of a tough insoluble film-forming resin such as a chlorinated rubber within which tributyltin fluoride isphysically dispersed12.When immersed in water, the freely dispersed molecules oftributyltin fluoride near the surface of the paint are able to diffuse out of the matrixof the paint film. As the biocide leaches out of the film, it leaves behind microscopicpores within the paint matrix. The inflow of sea water into these tiny pores causesthe release of fresh tributyltin biocide from beneath the surface layers of the film. Amajor disadvantage of this paint design is that with the passage of time, themicroscopic pores become clogged with insoluble materials thereby making it difficultfor biocide in the deeper strata of the paint matrix to be released. As a result,contact leaching antifouling paints work best only during the early part of their lifespan. When the antifouling action of the paint fails, a large amount of the biocideis still trapped in the inner matrix thereby creating a severe problem in the properdisposal of spent antifouling paints12.1.3.2 Ablative formulation.In this design, the tributyltin compound is dispersed into a film matrixcomposed of a mixture of soluble polymeric materials which are designed to breakdown over time12. As the film matrix breaks down, the biocide is released. Adisadvantage of this design is that it is difficult to control the actual breakdown ofthe paint film and the release of the biocide because the rate of paint film6breakdown is affected by water conditions and vessel speed4.1.3.3 Self polishing copolymer paints.In self polishing copolymer paints, the paint film is composed of a copolymerof methylmethacrylate and tributyltin methacrylate which also is the source of thebiocide. At the surface of the paint, sea water interacts with the hydrophobic copolymer and initiates a saponification reaction which cleaves tributyltin cation fromthe co-polymer backbone, releasing it into the sea. The release rate of the tributyltincation is gradual thereby enabling the biocidal action of the antifouling paint to lasta long time.1.4 TOXICiTY OF ORGANOTIN COMPOUNDS.In general, the toxicity of the organotin compounds RSnX4.. increases withthe increase in the number of alkyl or aryl substituents bonded directly to the tinatom15. Maximum toxicity is obtained when n=3. On increasing the number of alkylor aryl substituents above n=3, the toxicity drops. Therefore, tetraorganotincompounds on their own have no toxicity. The toxicity observed with thetetraorganotin compounds is believed to be due to their in vivo metabolism totriorganotin compounds13. As the number of carbon atoms in the alkyl chain isincreased above three, mammalian toxicity decreases.The type of R- group on the tin atom determines the level of toxicity tospecific organisms8.The trimethyltin compounds are the most toxic to insects while7the triethyltin compounds are the most toxic to mammals8. For gram-negativebacteria, the tripropyltin compounds are the most toxic8. The tri-n-butyltincompounds are the most toxic to gram-positive bacteria and fungi8.The toxic effect of the trialkyltin compounds is attributed to the inhibition ofmitochondrial oxidative phosphorylation, and subsequent disruption of a fundamentalenergy process13. The trialkyltin compounds bind to a number of proteins, andmortality may arise from direct reaction of the organotin species with proteins13.Differences in protein binding sites among groups of organisms would result in thevarying spectrum of effectiveness of the triorganotin compounds to differentorganisms’3’14Another mechanism by which trialkyltin compounds may derive their toxicityhas been described by Selwyn and Tosteson and Weith’6.According to theseauthors tributyltin’5”6,tripropyltin and triphenyltin’5 cations mediate chloride-hydroxide exchange in the mitochondria and smectic mesophases (liposomes)15,andin planar lipid bilayers16.Also, the tripropyltin cation has been reported to mediatechloride-chloride exchange across a lipid bilayer17. The ability of the trialkyltincations to mediate anion transport is the subject of study in chapter three of thisthesis.The dialkyltin compounds R2SnX also show a similar trend of decreasingtoxicity with increasing length of the alkyl chain8. The mode of toxicity of thedialkyltin compounds has been shown to be different from that of trialkyltincompounds18. The toxic action of the lower dialkyltin compounds is due to their8ability to combine with enzymes possessing two thiol groups in the correctconformation8’18 The biochemical effect of this is an interference with a-keto acidoxidation8’18The mono-organotin compounds RSnX3, do not show any important toxiceffect8.The organotin compounds do not appear to show any carcinogenic orteratogenic effect8. However, di- and tn- alkyl and aryl tin compounds have beenshown to induce chromosomal contraction in human lymphocytes19.Alterations in thespermatocyte chromosomes of the mesogastropod Truncatella subcvlindrica inducedby dibutyltin dichloride and tributyltin chloride have also been reported by Vitturiet a120, thus demonstrating the genotoxicity of these compounds.Of particular interest in the marine environment are the tributyltincompounds. Tributyltin compounds are very toxic to marine life at very lowconcentrations, and are suspected of inducing imposex in the female dogwhelkNucella lapillus21’2.Tributyltin compounds have also been reported to induce shellmalformations in the oyster Crassostrea gigas at very low concentrations’24,andhave also been reported to have caused high mortality in the larvae of the commonmussel Mytilus edulis25.The toxicity of tributyltin species to the following non-targetmarine organisms at the ppb level has been reported:- amphipod larvae26, lobsterlarvae and zoeal shore crab27, and the sheepshead minnow Cyprinodon variegatus28.At the low ppb levels, tributyltin species cause sublethal effects in the zoeal mud crabRhithropanopeus harnisii29 and copepods Acartia tonsa30.91.5 METABOLISM AN]) BEHAVIOUR OF TRIBUTYLTIN COMPOUNDSIN THE ENVIRONMENT.On introduction into the marine environment, tributyltin compounds areremoved from the water column by photolysis and assimilation by plants andanimals21,and by adsorption to the sediment, and particulate matter. Hydrolysis andvolatilization do not appear to be major degradative pathways22. The affinity oftributyltin compounds for sediments and particulate matter makes them far lessbioavailable to organisms in the upper water layer, but bottom feeding organisms areexposed to higher concentrations of tributyltin compounds. Therefore, the feedinghabit of a marine organism is an important factor in determining its tributyltin bodyburden. According to Maguire3’, tributyltin species adsorb so firmly to particles thatunder abiotic conditions, there was no desorption of tributyltin oxide from harboursediments over a period of ten months. However, under biotic conditions there wasmicrobial degradation resulting in the liberation of butylated and methylatedproducts. Tributyltin cation is hydrophobic, and therefore has a high tendency topreferentially accumulate in the surface microlayer of natural waters32. Its octanolwater partition coefficient (K0=55OO-7OO)33 and sediment-water partitioncoefficient(K0 = 16OO) values favour accumulation in the surface microlayer. Thesurface microlayer attracts and sequesters hydrophobic species such as tributyltinspecies. The preferential accumulation of tributyltin compounds in the surfacemicrolayer is expected to render tributyltin species unavailable to most organisms.However, bioaccumulation has been observed for a variety of organisms. Bacteria10and phytoplankton accumulate tributyltin species to concentrations 600 times and30,000 times respectively, greater than their exposure concentrations32’3.Also, abioaccumulation factor of 4400 has been reported by Evans and Laughlin36 for thehepatopancreas of the mud crab Rhithropanopeus harrisii. Accumulation oftributyltin species up to a concentration factor of 12,000 by the plant EelgrassZostera marina has been reported37.Tributyltin compounds have been found to exhibit preferential accumulationin certain tissues. Ward .j38 observed that the viscera of the sheepshead minnowcontained higher concentrations of tributyltin oxide than the cranial or muscle tissues.The reported bioaccumulation factors for tributyltin compounds are high enough towarrant concern with regard to their persistence and accumulation in food chains.However, they are degraded in vivo by bacteria39, fungi60, algae35 and fish34. Thedetoxification route for tributyltin compounds involves their conversion to the lesstoxic dibutyltin, monobutyltin, and inorganic tin species. Getzendaner and Corbin41have also reported similar detoxification pattern for tricyclohexyltin species. Barug4°has observed the degradation of tributyltin oxide to monobutyltin derivatives by thebacteria Pseudomonas aeruginosa. However, tributyltin chloride was not degradedunder anaerobic conditions by the same bacteria. Maguire have observed thein vivo degradation of tributyltin species by a green algae with the majordegradation product being dibutyltin species.Tributyltin compounds do not appear to be amenable to biomagnification.Macek ti42 have presented data indicating that chemicals with short or moderate11half-lives in vivo do not pose a biomagnification problem. Tributyltin species havea half-life considerably shorter than forty days by aerobic metabolism43.1.6 ANALYTICAL METHODS FOR ORGANOTIN COMPOUNDS.The early analytical methods available for the determination of tin wereclassical gravimetric or volumetric procedures that gave only total inorganic tinconcentrations. Beginning in the early 1930s, and extending into the early 1960s,optical spectrographic methods for total tin determination were extensively used forthe determination of geological samples. With the passage of time, the more sensitivecolorimetric, fluorimetric, neutron activation and flame atomic absorption techniqueswere introduced. The wide use of flame atomic absorption spectrometry washampered by the low sensitivity of the tin absorption lines44.For the organotin compounds, early analytical methods relied on theconversion of the organotin compounds to inorganic tin, usually by digestion withmineral acids followed by ignition. L.ater, more diversified techniques such aselectrochemical, chromatographic, and mass spectrometric techniques capable ofproviding speciation information were introduced. Recently, a tandem massspeetrometry (MS-MS) technique45 has been applied to a mixture of standardorganotin compounds with a view to analyzing them without prior derivatization tovolatile species or prior chromatographic separation. This method relies on the fixedrelationship between parent and daughter ions of any compound under fixedexperimental conditions. The applicability of this technique to the analysis of12environmental samples has not yet been demonstrated.Not much attention has been given to the qualitative analysis of organotincompounds. However, infrared spectroscopy46,Mossbauer spectrometry46’7,nuclearmagnetic resonance spectrometry48’95°and electron spin resonance spectrometry51have been applied to provide information on molecular structure and speciation.The various quantitative analytical methods applied over the years aredescribed in the following sections.1.6.1 Molecular spectrophotometry and spectrofluorimetric techniques.These methods rely on the attachment of a chromophoric ligand to theorganotin compound. This makes it possible to analyze the organotin compounds byusing uv-visible, or fluorescence spectroscopy, since the alkyl groups of the organotincompounds are not of much spectroscopic importance. A variety of ligands have beenemployed for this purpose. Aidridge and Cremer52 were the first to use dithizone forthe spectrophotometric determination of diethyltin and triethyltin species. Diethyltinand triethyltin chlorides react with dithizone to form colored complexes. Analysis ofthe complexes is effected following partitioning between aqueous potassiumhydroxide and chloroform. The diethyltin species partition into the alkali layer, whilethe triethyltin species migrate to the chloroform layer. The separated organotincompounds can then be determined by uv-spectrophotometry. A modified method forthe reaction between dithizone and the organotin compounds has been reported byHavir and Vretal53 for the determination of bis(tributyltin) oxide. Skeel and Bricker5413developed a spectrophotometric method for the determination of dibutyltindichioride by using diphenyl carbazone. This method achieved a sensitivity in themicrogram range. Other colorimetric reagents that have been used in the analysesof the organotin compounds are dithiol55, haematoxylin56, 8-hydroxyquinoline57,phenylfluorone58,pyrocathechol violet59’60,flavinol61 (3-hydroxyflavone).For the fluorimetric determination of the organotin compounds, 3-hydroxyflavone62and morin63 (2’ ,3 ,4,4’ ,5 ,7,-pentahydroxyflavone) have been appliedto the determination of phenyltin, and alkyltin compounds respectively.A major disadvantage of the spectrophotometric and spectrofluorimetrictechniques is the lack of specificity of these organic ligands to organotin compounds.However, limited selectivity can be achieved by employing various extractiontechniques prior to spectrophotometry or spectrofluorimetry.1.6.2 Electrochemical techniques.Electrochemical techniques rely on the difference in the redox potentials ofthe various organotin compounds for speciation. Polarography, in the various modessuch as anodic stripping voltammetry’65,and potentiometric titrations’67 havebeen used for the determination of organotin compounds in aqueous and non-aqueous media.Diethyltin dichioride was the first organotin compound whose polarographicreduction was recorded68.The polarographic behavior of other organotin compoundshas also been described69.The ease of the polarographic reduction of the organotin14compounds has been found to be a function of the alkyl moiety on the tin atom70.The ease of reduction follows the order7° ethyl > propyl > butyl. Polarography, inthe differential pulse mode has also been applied to the determination of organotincompounds71’273Potentiometric titration oforganotin compounds in dimethylsulfoxide (DMSO)has been reported to show well defined differential pulse polarographic peaks whoseheights are linearly dependent on concentration74. Quantitation of organotincompounds based on this observation has also been accomplished74.The half wave potentials of some organotin compounds have been determinedby Abeed et alTh, by using voltammetry, and cyclic voltammetry at rotating discelectrodes (gold and glassy carbon electrodes) in non aqueous solvents. Their resultsshowed that the reduction becomes more difficult as the electron donating ability ofthe alkyl or aryl groups attached to the tin atom increases (phenyl- methyl-buty1).The variation of the anion attached to the tin atom had little effect on the half wavepotentials. The application of electrochemical techniques to organotin analysis isrestricted by the sample matrix. Organic matter present in environmental samplescoats the electrode surface causing broadening of peaks, and shifts in peak potentials.1.6.3 Atomic spectrometry.Atomic absorption spectrometry has been used extensively in the analysis oforganotin compounds. Atomic absorption spectrometry is not capable of speciationunless a prior separation of the organotin compounds is achieved. The methods15usually employed to separate organotin compounds prior to their determination byatomic spectrometry include solvent-solvent extractions or conversion to organotinhydrides. The first analytical speciation and quantitation of organotin compounds byatomic absorption spectrometry after their derivatization to organotin hydrides wasdeveloped by Hedge et a176. The method involved the reaction of organotincompounds in natural water, acid digest of sediment or macroalgae with sodiumborohydride. The organotin hydrides produced were collected in a hydride trap whichwas cooled in liquid nitrogen. The hydride trap was warmed to release the organotinhydrides into a quartz tube furnace according to their boiling points. Since thepublication of this analytical method by the authors76, numerous atomic absorptionmethods based on hydride generation and the boiling point differences of theorganotin hydrides have been reported 77,78,79,80,81,82,83,84Many gas chromatographic techniques based on the hydride generationmethod of Hedge are presently in use (Section 1.6.4.1).The speciation of butyltin compounds by liquid-liquid extraction prior toatomic absorption spectrometry has been described by Mckie85. This method85involved the extraction of butyltin species into acidified hexane and the subsequentremoval of dibutyltin and monobutyltin species by washing with 3 % sodium hydroxidesolution. The hexane layer was evaporated and the solution of the residue in nitricacid was analyzed for tributyltin species by using graphite furnace-Atomic absorptionspectrometry (GFAAS). The matrix modifier used in this determination wasNH4H2PO.An analytical method similar to the technique reported by McKie85 has16been applied to the determination of tributyltin species in shellfish and sediments byStewart and de Mora. These authors employed K2CrO7as a matrix modifier.The use of atomic absorption spectrometry for the determination of organotincompounds is usually affected by severe matrix interferences. To overcome thisproblem, various matrix modifiers have been introduced. One of such matrixmodification methods was the coating of the interior surfaces of a graphite furnacetube with zirconyl acetate87. This has been shown to increase the atomizationefficiency of tin87. Peetre and Smith have reported that there is a relationshipbetween the atomic absorption sensitivity and the structure of an organotincompound. According to this report, the atomic absorption sensitivity decreased asthe energy of the alkyl-tin bond decreased.Atomic emission spectrometry of the organotin compounds has not beenpopularly used except in plasma emission89’9012 detection or flame photometricdetection in gas chromatography (Section 1.6.4.2a). Atomic emission spectrographicmethod has been described for the determination of bis(tributyltin) oxide93. Priorseparation of other organotin compounds present in the sample is necessary ifspeciation is desired because emission spectrography on itself, is not capable ofdistinguishing between the different organotin compounds.1.6.4 Gas chromatography.Organotin compounds are usually converted to volatile hydrides ortetraalkyltin compounds prior to gas chromatographic separations. In a few reports,17the analysis of organotin compounds without prior derivatization to hydrides ortetraalkyltin compounds has been accomplished94’56798.In such methods, thechromatograms are usually characterized by peak broadening and tailing. The columnefficiency is also decreased.Both packed and capillary gas chromatographic columns have been used withvarious detection techniques. The chromatographic techniques employed in theanalyses of the organotin compounds are discussed below.1.6.4.1 Hydride generation gas chromatography (HG-GC).This method involves the conversion of the organotin compounds to volatilehydrides by the use of excess borohydride to produce alkyltin hydrides of the formulaRSnH4..,. The generated hydrides are purged from solution with the help of an inertgas, and can be cryoscopically trapped. The cold trap is subsequently warmed, torelease the organotin hydrides into the column of the gas chromatograph. Detectionof the separated organotin hydrides is achieved by using various gas chromatographicdetectors.Woollins and Cullen99 have described a hydride generation-GC flameionization detection technique based on the method previously developed by Hodge76 for the analysis of organotin compounds. Hattori LV°° have also describedthe determination of organotin compounds in environmental water and sediments,on a packed column by using HG-GC electron capture detection. An ultratracemethod for the analysis of aquatic butyltin by HG-GC with flame photometric18detection has been described by Matthias et a1101. A novel on-column hydridegeneration analysis of organotin compounds by gas chromatography-atomicabsorption spectrometry (GC-AAS) has been described by Clark Li102,and TakamiJ103.In the method described by Takaini t.i103,fish samples were extracted withhydrochloric acid-ethanol mixture. The extracted organotin compounds in the fishwere transferred to ethylacetate/hexane by using liquid-liquid extraction, and thenapplied onto a sulfonated cation exchange column where the organotin compoundswere trapped. On-column hydride generation was effected by passage of an ethanolicsodium borohydride solution through the cation exchange resin. The generatedhydrides were extracted into hexane, and analyzed by GC-MS.The GC-hydride generation method of organotin determination has been veryextensively used by numerous authors104’5’°678”9.Two detection methods have widely been used in gas chromatography for theanalyses of organotin compounds after their conversion to volatile derivatives. Theseare flame photometric detection (FPD) and atomic absorption spectrometry (AAS).Coupling the gas chromatograph directly to an atomic absorption spectrometer (GCAAS) appears to be the most popular technique for element specific speciation ofthe organotin compounds after hydridization.Methyltin species in natural water have been determined after hydridederivatization by using GC-graphite furnace atomic absorption spectrometry (GCGFAAS)’10.Butyltin species in natural water and sediments have been analyzed byQuevauviller and Donard’1’, by using GC-HG-quartz tube atomic absorption19spectrometry.In enviromnental samples, production of volatile hydrides may be inhibited bysevere matrix interferences112• Such matrix interferences can be eliminated by the useof L-Cysteine”3.1.6.4.2 Conversion to telraalkyltin compounds.Conversion of organotin compounds to tetraalkyltin derivatives is usuallyaccomplished by reacting the organotin compounds with a Grignard reagent, orsodium tetraethylborate. The reaction of monoalkyltin, dialkyltin, and trialkyltinspecies with the Grignard reagent proceeds to completion at very low concentrations,and no rearrangement of the original alkyl groups attached to the tin atom is usuallyrved14 The tetraalkyltin derivatives formed are usually stable in organicsolvents114’5.Various alkyl groups such as methyl116’7,ethyV’8,pentyV19”20,andn-hexyV21.122, have been attached to the butyltin compounds to facilitate theiranalyses. Detection of the derivatized tetraalkyltin compounds is usuallyaccomplished by the use of various gas chromatographic detectors. The GC-detectorsystems that have been used for tetraalkyltin analyses are described below.1.6.4.2(a) Gas chromatography with flame photometric detection (GC-FPD).The flame photometric detector has been used extensively for the detectionand quantitation of organotin compounds in the environment following theirderivatization to tetraalkyltin compounds. Developed by Aue and Flinn’, flame20photometric detection has been used for tin-specific detection in gas chromatographicanalyses of butyltin species infishUS,lZO,waterl2Ol24,sediments120’’225,andfor the determination of methyltin species in water126.A disadvantage of the flame photometric detector is the decrease in sensitivitywhich may be caused by the accumulation of Sn02 on the internal surfaces of thedetector, and by tropolone, a ligand sometimes used in the extraction of organotincompounds127•Flame photometric detection of the organotin compounds relies on theconversion of tin to Snil in air/hydrogen flame. SnH yields a red emission line in thegas phase at about 610 nm and almost all analyses for tin have been carried out atthis wavelength. Earlier, Aue and jjflfl23 had described another emission line fortin at 390 nm in the flame photometer.This emission line was unstable, irreproducibleand easily quenched123, and was later attributed to a quartz surface induced tinluminescence’28.Jang Ii29 have described a sensitive flame photometric analysisinvolving the 390 nm emission line. According to the authors, a clean quartz surfaceis required in the vicinity of the flame to achieve stability of the emission line, andonly the Shimadzu flame photometric detector has this feature. The detection limitwas claimed to be about thirty times better than that at the 610 nm emission line.1.6.4.2(b) Gas chromatography with atomic absorption spectromeiric detection(GC-AAS).Analysis of organotin compounds by GC-AAS after conversion to tetraalkyltin21compounds is not popular and has only been seldomly used. Forsyth et a113° haveapplied this method to the determination of organotin compounds in fruit juices.1.6.4.2(c) Gas chromatography with mass spectromeiric detection (GC-MS).GC-MS affords a very reliable method in the analysis of the organotincompounds, since detection is based both on retention data and fragmentationpattern. The first application of GC-MS to the analysis of organotin compounds wasdescribed by Meinema In this procedure, benzene extracts of pure butyltincompounds were converted to their butylmethyltin derivatives by usingmethylmagnesium bromide. The derivatized butylmethyltin compounds were analyzedby using GC-MS with dibutyihexylmethyltin as the internal standard. The applicationof GC-MS to the determination of organotin compounds in environmental sampleshas been reported by Cullen etalm, Forsyth.Lai’30,Muller131, Unger t.i132.1.6.5 Liquid chromatography (LC).Liquid chromatography of the organotin compounds does not requirederivatization to volatile species, and hence could be useful for the analysis of nonvolatile organotin compounds. The alkyltin species are difficult to detect by uv-visiblespectroscopy therefore, derivatization might be necessary to enhance their detection.Various liquid chromatographic detectors have been used for the analysis oforganotin compounds. An indirect photometric method for the determination ofalkyltin compounds has been described by Whang t.ilV33 for tributyltin, tripropyltin,22triethyltin, and trimethyltin species after their separation on a strong cation exchangecolumn.For high sensitivity, inductively coupled plasma-mass spectrometers (ICPMS)’34’135”6,inductively coupled plasma-atomic emission spectrometers (ICPAE)135”36 and atomic absorption spectrometers137’89401 have been used todetect organotin compounds in high performance liquid chromatography (HPLC).Direct coupling of the liquid chromatograph to a mass spectrometer or atomicabsorption spectrometer is associated with problems such as solvent interferences.The large amount of solvent and sometimes non-volatile buffers that go into thedetector systems are also a major concern. This concern can be addressed byinterfacing the LC and the detector systems, or by the use of a microbore column.The small solvent flow rate (10-100 iL/min) in microbore HPLC has been shown tobe compatible with direct effluent introduction to a flame atomic absorptionspectrophotometer’42.A coupled LC-AAS technique with enhanced laser ionization detection hasbeen described by Epler et al’43, for the analysis of tributyltin species in sediment.In their technique, tributyltin species in a sediment sample were extracted into 1-butanol, separated on a strong cation exchange column and detected by using apremixed air-acetylene flame which was irradiated with two pulsed lasers. Theenhanced ionization of the tin atom in their technique143 was attributed to a rapidcollisional ionization of the excited tin atoms in the flame.Another method of solving the problem of excessive solvent introduction into23the detector system, is by the post column derivatization of the organotin compoundsto volatile hydrides which can then be introduced into an atomic absorptionspectrometer144. This method has been applied to the determination of methyltinspecies144.Organotin compounds adsorb strongly onto unmodified silica gel columnstherefore, most liquid chromatographic separations are performed on reversed phasecolumns, size exclusion columns, and other modified columns such as cyano bondedsilica gel columns. Jessen et al145 have studied the adsorption behavior of alkyltinhalides on various chromatographic columns, and concluded that silica basedoctadecyl (ODS) and cyano columns are not sufficiently inert to alkyltin halides whilesilica gel columns pyrolytically coated with carbon black are inert. However, theorganotin halides have been reported to adsorb on a commercially availablegraphitized carbon black146.The ability of organotin halides to adsorb on graphitizedcarbon black was the basis for a selective determination of the organotin compoundsby Fern et al’46.The separation of the alkyltin halides on a cyanopropyl bonded silica gelcolumn and their detection by fluorescence spectrometry after on-columncomplexation with morin has been described by Langseth’47.It has been reported byJessen et al’45, that rearrangement of the alkyl groups on the tin atom can occurespecially if tetraalkyltin compounds are co-injected with other organotin compoundsinto the LC column.241.6.6 Thin layer chromatography (TLC) and high performance thin layerchromatography (HPTLC).Thin layer chromatography is not widely used for the quantitative analysis ofthe organotin compounds. However, it has been applied to qualitatively identifyorganotin compounds. The separation of butyltin compounds and their colorimetricdetection after complexation with pyrocatechol violet has been reported by Laughlinet a1148.Speciation of the mammalian organotin metabolic products has been reportedby Kimmel et a1149. Their method involved both normal and two dimensional TLCtechniques, and visual detection of the organotin species after complexation withcolorimetric reagents such as pyrocathechol violet or dithizone. Vasundhara Li56have demonstrated the good resolving power of the TLC for a series of tn- and diorganotin compounds. Detection of the separated organotin compounds was bytreatment of the TLC plate with haematoxylin.High performance thin layer chromatography (HPTLC) has not beenpopularly used for qualitative analyses of the organotin compounds. However thequantitation of butyltin compounds in a wood matrix, by using HPTLC has beenreported by Ohlsson ct.i150. Their method involved the post column developmentphotolysis and complexation of the butyltin species with pyrocatechol violet, followedby colorimetry of the tin-pyrocatechol violet complex.A HPTLC method of quantitation has also been described by Tomboulian2j151 for phenyltin compounds. Quantitation was by fluorescence scanningS 25densitometry, following in situ complexation of the phenyltin compounds with morin.1.7 TRIBUTYLTIN AND GOVERNMENT REGULATIONS.Tributyltin compounds dissolved in marine waters exhibit acute toxicity to avariety of aquatic life. Available data in the literature tend to suggest that fish andlarger crustacea are less sensitive to tributyltin compounds than bivalves, molluscs,phytoplankton and small crustaceans. It has been established that molluscs aregenerally very sensitive to organotin compounds152.Following the establishment ofa correlation between tributyltin compounds, shell malformation and abnormalgrowth in oysters, the French Government in 1982 banned the use of antifoulingpaints containing more than 3 % by weight of tributyltin compound on boats less than25 tons. In 1987, a total ban on the use of organotin paints on vessels less than 25metres came into effect in France’52.In 1986, the Government of England prohibited the retail sale of someantifouling paints containing tributyltin compounds. In 1987, the use of tributyltincontaining paints on small boats and mariculture equipment was banne&53.The use of organotin compounds in fresh water antifouling paints is prohibitedin Germany and Switzerland’54.In Canada, tributyltin compounds are registered under the Pest ControlProducts Act for use as a slimicide and for general lumber preservation. Its use asa preservative for nets is not allowed”. In 1989, the Canadian Governmentprohibited the use of tributyltin compounds on vessels less than 25 metres in length,26and also stipulated a maximum release rate of 4 j.tg tributyltin compound per squarecentimetre of ship’s hull155.1.8 OBJECTIVES AND SCOPE OF THE PRESENT STUDY.Tributyltin compounds have been shown to cause shell malformation in oyster,and other molluscs11’234,and have also been linked to imposex in the marinegastropods such as the female dogwhelk10.Cullen Li”7have reported the presenceof butyltin and cyclohexyltin species in some coastal areas of British Columbia,Canada. The toxic effect of the cyclohexyltin species is not known with certainty.Therefore, it is necessary to provide data on the extent of organotin pollution in thecoastal areas of British Columbia, Canada.Chapter 2 of this thesis provides information on the levels of butyltin anddicyclohexyltin species in some marine locations of British Columbia.Although the trialkyltin compounds are thought to exert their toxicity by theinhibition of mitochondrial oxidative phosphorylation, little attention has been givento the effect of the alkyltin compounds on biomembranes. Early studies by Selwyn15,and Tosteson and Wieth’6 showed that tributyltin, triphenyltin cations act as carriersfor CF and OW, and therefore mediate CF/OW, while the propyltin cationmediates CF/CF exchange across artificial biomembranes called liposomes. Later,Tosteson and Weith156 showed that tributyltin chloride affects the dipole potentialof phosphatidylethanolamine lipid bilayer, and lowers it by about 70 millivolts. Theeffect of organotin compounds on the other properties of membranes such as27permeability, elasticity, etc has not been reported. A knowledge of the effect of theorganotin compounds on the other properties of the membrane is important for acomplete understanding of the mechanisms of their toxicity.Arakawa et a1157 have reported the inhibition of intracellular calciummobilization by tributyltin chloride and dibutyltin dichioride. Arakawa and Wada’58then surmised that the inhibition of Ca2 mobilization may be due to changes in themembrane structure caused by the butyltin compounds. Also, the authors158 hadsuggested that the toxicity of the alkyltin compounds should depend on theirsolubility in biological fluids, and their extent of incorporation into cells.Therefore, the experiments described in Chapter 3 of this thesis are concernedwith providing information on the permeability changes of model biomembranesknown as liposomes, and the effect of organotin incorporation into these liposomeson membrane permeability.It is necessary to use liposomes as models for biological membranes becauseof their similarity with true biomembranes (Chapter 3 Section 3.3). Also, the use ofliposomes eliminates complications that may arise in interpreting experimental dataif true biomembranes are used for permeability studies.The ease or difficulty of efflux of an encapsulated probe permeant;dimethylarsinic acid (DMA) from these liposomes in the presence and absence of theorganotin compounds, is an indication of how the membrane permeability respondsto the presence of the organotin compounds.Since the advent of organotin pollution in the marine environment, every28effort by workers in the field of environmental pollution has been directed towardsproviding data on the level and speciation of the organotin compounds. The total tincontent of marine animals has been largely neglected. Therefore, Chapter 4 of thisthesis is concerned with total tin determination in oysters, and the analytical methoddevelopment for total tin determination by hydride generation-atomic absorptionspectrophotometry.29CHAPTER 2SPECIATION AND QUANT1TATION OF BUTYLTIN ANDCYCLOHEXYLTIN COMPOUNDS IN MARINE ORGANISMS BY USINGCAPILLARY COLUMN GC-MS SIM.2.1 INTRODUCTION.The work reported in this section involved the analysis of marine animals fororganotin compounds, and the synthesis of standard tetraorganotin compounds whichwere used as calibration standards for quantitation. The speciation and quantitationprocedure involved a prior extraction of the organotin compounds from the marineorganisms by the use of methylene chloride. The extracted organotin compoundswere reconstituted in n-hexane and subjected to Grignard methylation, a well knownchemical reaction to yield tetraorganotin compounds. Speciation and quantitationwere by capillary column GC-MS SIM.The high sensitivity and specificity of the mass spectrometric detectorespecially for tin compounds makes it the detector of choice. Tin has thirteen stableisotopes which in mass spectrometry give rise to a very characteristic isotope pattern.The isotope pattern for tin is very diagnostic for distinguishing tin compounds fromother compounds that may co-elute with them during chromatographic separations.The use of the mass spectrometer as a detector for gas chromatographic separationsis accomplished by the direct coupling of the GC capillary column to the ion sourceof the mass spectrometer.302.2 EXPERIMENTAL2.2.1 Instrumentation.2.2.1.1 Gas chromatography (GC).The gas chromatograph used to establish the retention times, and theoptimum chromatographic conditions for organotin separation was a Hewlett PackardModel 5890 instrument equipped with a flame ionization detector (FID). Dataacquisition from the gas chromatograph was achieved by using a Hewlett Packard3393A integrator. The GC column was a DB-1 polysiloxane stationary phase wallcoated open tubular (WCOT) capillary column (15 m x 0.25 mm i.d) purchased fromJ & W Scientific, Folsom, California. The carrier gas was helium at a linear velocityof 30 cm/s. The column temperature was held at 50°C for 10 minutes, and thenincreased at the rate of 20°C per minute to a final temperature of 240°C untilcomplete elution was obtained.2.2.1.2 NMR and mass spectrometry.1H NMR spectra were run at 300 MHz by using a Varian XL 300spectrometer. Chemical shifts are quoted relative to tetramethylsilane as externalreference. 119Sn NMR spectra were obtained on the same instrument. 119Sn chemicalshifts are quoted relative to tetramethyltin as external reference.Low resolution mass spectra (using electron ionization, El) for characterizing31the calibration standards were obtained on a Kratos MS 50 mass spectrometer.2.2.1.3 Gas chromatography-mass speciromeiry (GC-MS).The GC-MS system consisted of a Carlo-Erba Fractovap series 4160 gaschromatograph interfaced to a Kratos MS 80 RFA double focusing massspectrometer equipped with a Kratos DS55 data system. The mass spectrometer wasoperated in El under selected ion monitoring mode (SIM), and was calibrated byintroducing perfluorokerosine into the ion source. The operating conditions for themass spectrometer were; calibration range = 118-331, sweep = 1500 ppm, cycle time= 1 second, filament current = 1-2 Amp., electron voltage = 70 eV., ion sourcetemperature = 180 °C.The gas chromatograph was operated in the temperature programming mode.The temperature of the column was kept constant at 50 °C for 10 minutes, and thenincreased to 240 °C at a rate of 20 °C per minute until complete elution wasobtained. The injector temperature was maintained at 250 °C. The GC column is asdescribed in section 2.2.1.1, and was connected to the mass spectrometer via acapillary interface. The carrier gas was helium at a linear velocity of 30 cm/s.2.2.1.4 Mechanical shaker and blender.The mechanical shaker employed during the extraction step of the organotincompounds from environmental samples was a Magniwhirl reciprocating shaker BlueM Electric Company, Blue Island, Illinois, U.S.A.).32The blender used to homogenize the biological samples was obtainedcommercially.2.2.2 Materials and reagents.Dibutyltin dichioride and tributyltin chloride were purchased from M & TChemicals Inc., Rahway, New Jersey, and Ventron (Alfa Inorganics) BeverlyMassachusetts, U.S.A. respectively. Tin metal (20 mesh) was obtained fromMallinckrodt Chemical Works, St Louis, Missouri, U.S.A.. Tricyclohexyltin chloride(Technical Grade) and methylmagnesium bromide (3M in diethyl ether) wereobtained from Aldrich Chemical Company, Milwaukee, U.S.A.. Dicyclohexyltindibromide was purchased from Johnson Matthey (Alfa products), Danvers,Massachusetts, U.S.A. .The following chemicals silica gel (230-400 mesh) and sodiumchloride (Reagent Grade) were purchased from BDH, Poole, England. lodobutaned9 and 2-ethoxyethanol were supplied by Merck Frosst Canada Inc. (MSD isotopedivision) Montreal, Canada and Matheson, Coleman and Bell. ManufacturingChemists, Norwood, Ohio, U.S.A. respectively. The following reagents and solventswere procured from Fisher Scientific, Fair Lawn, New Jersey, U.S.A :- methyl iodide(Certified Grade), anhydrous magnesium sulfate (Certified Grade), anhydrous diethylether (Reagent Grade), n-hexane (HPLC Grade), n-pentane (Spectrograde), nheptane (HPLC Grade).332.2.3 Synthesis of standard organotin compounds.2.2.3.1 Synthesis of tributylmethyltin, (C4H9)3SnCH.Tributyltin chloride (2.99 g, 0.0092 mol) was dissolved in n-hexane (100 mL)in a 250 mL Erlenmeyer flask. Excess methylmagnesium bromide (6 mL of 3M,0.0 180 mol) in ether was added to the reaction mixture and stirred for six hours atroom temperature, after which the excess methylmagnesium bromide was destroyedby the gradual addition of sulphuric acid (1 M) while the reaction flask was cooledin an ice bath. The reaction mixture was transferred to a separatory funnel where theaqueous layer was removed, and the hexane layer was washed five times withhydrochloric acid (10 mL of 10% HC1), dried with anhydrous sodium sulphate andfiltered into a round bottom flask (250 mL). The hexane was removed on a rotaryevaporator to yield the crude product which was distilled twice at reduced pressureto obtain 1.70 g (61% yield) tributylmethyltin, b.p 58°C I 10 mm Hg. Analysis: %Found: C, 51.40; H, 9.80. % Calcd: C, 51.18; H, 9.91.2.2.3.2 Synthesis of dibutyldimethyltin (C4H9)2Sn(CH3.Dibutyltin dichloride (1.46g, 0.0048 mol) and excess methylmagnesiumbromide (15 mL of 0.045 mol) were reacted for six hours as described in Section2.2.3.1 to yield a crude product which was distilled twice to obtain 0.9 g ofdibutyldimethyltin (71% yield), b.p 33°C I 4 mm Hg. Analysis: % Found: C, 45.83;H, 9.30. % Calcd: C, 45.67; H, 9.20.342.2.3.3 Synthesis of tricyclohexyimethyltin (C6H11)3SnCH.Tricyclohexyltin chloride (2.02g. 0.0050 mol) and excess methylmagnesiumbromide (10 mL of 3M, 0.030 mol) were reacted for 12 hours as described in Section2.2.3.1. The crude product was distilled to obtain 1 g tricyclohexylmethyltin (52%yield), b.p 128°C / 0.6 mm Hg. Analysis: % Found: C, 59.93; H, 9.47. % Calcd: C,59.56; H, 9.47.2.2.3.4 Synthesis of dicyclohexyldimethyltin (C6H11)2Sn(CH3.Dicyclohexyltin dibromide (2.00g. 0.0045 mol) and excess methylmagnesiumbromide (10 mL of 3M, 0.030 mol) were reacted for 12 hours as described in Section2.2.3.1. The obtained crude product was distilled to obtain 0.87gdicyclohexyldimethyltin (61 % yield), b.p 88°C / 0.06mm Hg. which was identified bymass spectrometry and NMR spectrometry (Section 2.4.1).2.2.3.5 Synthesis of the internal standards.2.2.3.5(a) Synthesis of tetrapropyltin (C3H7)4Sn.Tetrapropyltin was synthesized according to the method described fortetraethyltin159,by reacting magnesium turnings (8.26 g, 0.34 mol), n-propyl bromide(46.74 g, 0.38 mol) and tin(IV) chloride (13.81 g, 0.053 mol) in anhydrous diethylether. The obtained crude product was distilled twice to yield 8.5 g tetrapropyltin, b.p108°C / 11 mm Hg. Analysis: % Found: C, 49.61; H, 9.76. % Calcd: C, 49.52; H,359.70.2.2.3.5(b) Direct synthesis of(C42H9)SnI and (C42H9)3SnI and the subsequentsynthesis of deuteriated internal standards (C421L)3SnCH and(C42H9)Sn(CH3.This synthesis was carried out according to the method reported by Oakes andHutton16° for dibutyltin diiodide. Deuteriated butyliodide (3. 13g, 0.017 mol) and 2-ethoxyethanol (1 mL) were mixed together in a round bottom flask (50 mL). Lithium(0.097g, 0.014 mol) and tin metal (0.89g, 0.0075 mol) were also added to the roundbottom flask. The contents of the flask were refluxed for two hours with stirring, andvacuum distilled to obtain 0.77 g of a mixture of the crude products (C42H9)SnI(71% yield) and (C42H9)3SnI (29% yield) as identified by GC-MS. The crudeproducts in pentane were treated with aqueous potassium hydroxide (2% w/v) toprecipitate the deuteriated dibutyltin dihydroxide (mlz=286). The pentane solutionwas filtered and evaporated under reduced pressure to give deuteriated tributyltinhydroxide as identified by mass spectrometry (mlz=335).Aliquots (0.18 g) of the deuteriated tributyltin. hydroxide and dibutyltindihydroxide were each reacted with methylmagnesium bromide (0.3 mL, 0.0009 mol)in hexane as described in Section 2.2.3.1 to give (C42H9)3SnCH and(C42H9)Sn(CH3 respectively as identified by GC-MS.362.2.3.6 Synthesis of methylmagnesium iodide.Magnesium turnings in slight excess (O.062g, 1.50 mol) and methyl iodide(210.07g, 1.48 mol) were reacted in anhydrous diethyl ether according to standardprocedure161 to obtain the Grignard reagent, methylmagnesium iodide.2.3 ANALYTICAL PROCEDURE.2.3.1 Gas chromatography.2.3.1.1 Establishment of elution profile and retention data.Appropriate amounts of the well characterized standard tetraorganotincompounds (C4H9)3SnCH,(C49)2Sn(CH3,(C6H11)3SnCH,(C6H11)2Sn(CH3 and(C3H7)4Sn were each dissolved in n-heptane in different volumetric flasks (25 mL)to form the stock solutions (50 tg/mL as Sn). A working solution (5 ig/mL as Sn)of each tetraorganotin compound was prepared in n-heptane from the stock solutions.Aliquots of each working solution (1 J.LL) of each standard tetraorganotinsolution (5 gImL as Sn) were injected into the capillary column of the gaschromatograph by using splitless injection. The retention time of each standardorganotin compound was noted.Aliquots of each stock solution were pipetted into the same volumetric flask(5 mL) to form a mixture of all the standard tetraorganotin compounds (5 jgImLas Sn). This mixture (1 L) was separated on the capillary column of the gas37chromatograph. Each of the tetraorganotin compounds was detected and identifiedon the basis of the earlier established retention times.2.3.1.2 Suitability of tetrapropyltin as internal standard.The suitability of tetrapropyltin as internal standard was verified on thetributylmethyltin solutions according to the following procedure:-four sets oftributylmethyltin solutions in heptane were prepared. Each set of solutions contained2, 4, 6, 8, 10 g/mL (as Sn) tributylmethyltin, and also contained the internalstandard tetrapropyltin at only one of the following concentration levels:- 2 ,4,6,50g/mL as Sn. Each mixture of tributylmethyltin and the internal standard (1 L) wasinjected into the capillary column of the gas chromatograph and separated. Threereplicate injections of each sample solution were made. The peak area ratios of thetributylmethyltin to the internal standard were plotted against the variousconcentrations of the tributylmethyltin solutions (Section 2.4.4).2.3.1.3 Suitability of (C42H9)3SnCH and (C42H)Sn(CH3) as internalstandards.A mixture of the deuterated compound (C42H9)3SnCH,and tributylmethyltinin heptane was injected into the capillary column of the gas chromatograph. Noseparation was obtained. Similarly, no separation was obtained for a mixture of(C42H9)Sn(CH3 and dibutyldimethyltin. Although no separation was obtained onthe gas chromatographic column, the deuterated butyltin compounds can still be used38as internal standards if high instrument sensitivity is not desired, because underconditions of GC-MS SIM, the butyltin compounds and their analogous deuteratedcompounds co-elute, and are simultaneously detected by the mass spectrometer.Therefore at a given retention time, only a few number of scans can be obtained foreach fragment ion, thereby leading to decreased sensitivity.2.3.2 Low resolution mass spectrometry: selection of fragment ions used forselected ion monitoring CC-MS.The tetraorganotin compounds, as neat liquids were subjected to lowresolution mass spectrometry. The fragment ions, and their intensities were recorded(Section 2.4.2 Tables 2.2 to 2.4). Fragment ions possessing the highest intensitieswere selected to be monitored in the GC-MS SIM analysis.2.3.3 GC-MS retention data, calibration curves, and precision.Appropriate amounts of the well characterized standard tetraorganotincompounds, (C4H9)3SnCH(C49)2Sn(CH3,(C6H11)2Sn(CH3 and (C6H11)3SnCHwere each dissolved in n-heptane in separate volumetric flasks (25 mL) to formstandard solutions. Appropriate amounts of each standard solution were pipetted intothe same volumetric flask (50 mL), and made up to the mark with n-heptane to formthe stock solution (5 tgImL as Sn, of each of the standard tetraorganotincompounds). Appropriate amounts of the stock solution were placed into six differentvolumetric flasks (10 mL) together with aliquots of the tetrapropyltin solution in39heptane, so that each volumetric flask contained 0.2 g/mL (as Sn) of the internalstandard, and 0.2 to 1.2 tg/mL of a mixture of all the tetraorganotin standards.Each mixture of the standard organotin compounds and the internal standard(1 jL) was analyzed by GC-MS SIM. Only the fragment ions shown in Table 2.7(Section 2.4.3) were monitored at the retention times shown in Table 2.6. The peakarea ratios of the standard organotin compounds to the internal standard wereplotted against the standard tetraorganotin concentrations (Section 2.4.5, Figures 2.8and 2.9) to afford the calibration curves.2.3.4 Recovery studies.Recovery studies were performed in duplicate at one level of organotinconcentration (1.5 tg as Sn). Shrimp (Pandalus tridens) (40 g, wet wt) from a batchwhose prior analysis by GC-MS revealed the absence of any organotin compound washomogenized in a blender, and transferred to an Erlenmeyer flask (1 L). The shrimphomogenate was spiked with 1.5 g (as Sn) of each of the following compounds:tributyltin chloride, dibutyltin dichioride, Iricyclohexyltin chloride, and dicyclohexyltindibromide all dissolved in n-heptane. The solution was vortex mixed and sodiumchloride (20 g) was added together with concentrated hydrochloric acid (50 mL) andmethylene chloride (100 mL). The resulting slurry was shaken on a mechanicalshaker for one hour and filtered through a pyrex glass wool into a separatory funnel(1 L) where the lower methylene chloride layer was removed. The shrimp residue,and the glass wool were returned to the Erlenmeyer flask (1 L). The aqueous layer40from the separatory funnel was also returned to the Erlenmeyer flask (1 L) and theextraction procedure repeated twice more. All the methylene chloride layers werepooled together, dried with anhydrous magnesium sulfate and filtered through aWhatman No 1 filter paper into a round bottom flask (500 mL), and evaporated offby using a rotary evaporator to obtain an oily residue which was dissolved in nhexane solution.Aliquots of the shrimp extract in n-hexane (5 mL), and a solution of theinternal standard (0.5 mL of 2.2 .tgImL as Sn) and methylmagnesium iodide indiethyl ether (3 mL) were added to the Erlenmeyer flask. The reaction mixture wasleft standing with intermittent shaking for one hour, after which excessmethylmagnesium iodide was destroyed by the gradual addition of dilute sulphuricacid (1 M) and de-ionized water (about 10 mL). The hexane solution in theErlenmeyer flask was transferred to a separatory funnel (50 mL) where the aqueousbottom layer was removed, and the upper hexane layer passed through a silica gelcolumn (2 cm x 0.5cm i.d) pre-equilibrated with n-pentane. The hexane solution wasdrained down the silica gel column, and then eluted with n-pentane (15 mL) into asample vial. The solution in the sample vial was concentrated down to about 0.2 mLby blowing a gentle stream of nitrogen. About 0.3 mL of n-heptane were added tothe sample vial, to bring the total volume of the solution to about 0.5 mL. Theheptane solution (1 tL) was analyzed in duplicate by GC-MS SIM.412.3.5 Extraction of the organotin compounds from marine animals.About 2-5 frozen marine bivalves were thawed, and then shucked. Portions ofthe soft tissue weighing between 18-137 g were placed in a blender together with 100mL de-ionized water, and homogenized for about 3 minutes. The homogenized tissueslurry was transferred to an Erlenmeyer flask (1 L). Sodium chloride (20 g),concentrated hydrochloric acid (50 mL), and methylene chloride (100 mL) wereadded into the tissue slurry and mixed by shaking. The tissue slurry was then shakenon a mechanical shaker for one hour, and then extracted and processed as describedin Section 2.3.4 above. A flow diagram for the extraction of organotin compoundsfrom marine animals is given in Figure 2.1.42Filter through glass woolTransfer into separatory funnelRemove CE2CI2 layerRepeat CH2CI2_____________________extraction twiceRemove CH2CIZRe-dissolve In u-HexaneDerivatize to tetraalkyltinSilica gelclean upTissueIHomogenizelOOmL deionized water5OmL MCi20g NaCIlOOmL CR2C1ZShake for lhrPool CHzC1Z layer CR2 C12layerInject into GC-MSFigure 2.1 Flow diagram for the extraction of organotin from marine animals.432.4 RESULTS AND DISCUSSION.2.4.1 Characterization of the standard tetraorganotin compounds.The standard tetraorganotin compounds were synthesized by Grignardmethylation of butyltin and cyclohexyltin halides as described in Section 2.2.3. above.The butylmethyltin and the tetrapropyltin compounds afford good elemental analyses.Tricyclohexylmethyltin was characterized by elemental analysis and massspectrometry (Figure 2.2). Dicyclohexyldimethyltin was characterized by 1H NMR(Figure 2.3) and mass speetrometry (Figure 2.4). The integrated peak area ratios ofthe methyl group protons, and the cyclohexyl group protons in the 1H NMR spectraof dicyclohexyldimethyltin were used to confirm the number of protons on thecyclohexyl group. The number of protons on the cyclohexyl group ofdicyclohexyldimethyltin was 22 as expected. In the NMR spectra of the freecyclohexane162,all the protons are equivalent, and give rise to a singlet at O = 1.4ppmin CC!4.In substituted cyclohexane such as bromocyclohexane, two groups of protonsare observed162. In dicyclohexyldimethyltin three groups of cyclohexyl protons areobserved. The peaks are strongly coupled, and the multiplets observed cannot beexplained on the basis of a first order coupling. The cyclohexyl group proton on thecarbon atom directly bonded to the tin atom is deshielded and resonates at low field(o=1.8 ppm).Further characterization of the butylmethyltin and cyclohexylmethyltincompounds was provided by 9Sn NMR (Table 2.1) as this information is not yet44available in the literature. From the chemical shift data shown in Table 2.1, it canbe inferred that the cyclohexyl group has a more deshielding effect on the tin atom,than the n-butyl group.Table 2.1 119Sn NMR chemical shifts for the slandard organotin compound?.Compound 6 (ppm) Conc. (M)(C4H9)3SnCH -6.128 1.10(C4H9)2Sn(CH3 -2.539 0.58(C6H1,)3SnCH -44.378 0.77(C6H,,)2Sn(CH3) -16.393 0.30(C3H7)4Sn -18.072 1.98a=119Sn NMR was obtained on deuterated chloroform solutions of theorganotin compounds, and referenced relative to tetramethyltin.45++c-)L)C%1‘1-—‘0‘0 L.)c_)100 219+c-)13550 +L)z4.‘I)z? ‘0I L0 mi TT384100 150 200 250 300 350 400 450 500MJZFigure 2.2 Mass spectra (El) of iricyclohexylmethyltm.46Figure 2.30PPM6 5 4 3 2 11H NMR spectra of dicyclohexyldimethyltm.47+N,‘0L)+N,c-)C#•)100Cl)zz0+N,C,,+N,c-)‘0L)++r.jN,c-)Cl)rj‘0L)+N,ci)C%J‘0L)+C,)‘0C.)100 150 200 250 300MJZ350Figure 2.4 Mass spectra (El) of dicyclohexyldimethyltm.482.4.2 Fragment ions and intensities of the standard tetraorganotin compounds.The standard tetraorganotin compounds were subjected to low resolution massspectrometry. The molecular ions were of low intensity, and therefore were not usedfor the GC-MS analysis. The fragmentation involves the loss of butyl or cyclohexylor methyl groups from the tin atom. The fragment ions and intensities relative to thebase peak are given in Tables 2.2, 2.3 and 2.4.Table 2.2 Major fragment ions of tributylmethyltin.Standard M/Z Fragment ion Relative Intensitycompound (% )(C4H9)3SnCH 121 SnH4 79(MW=306)8 135 CH3Sn 100177 C4H9Sn 65193 C4H9SnHCH3 99235 (C4H9)2Sn 12249 (C4H9)2SnCH3 93291 (C4H9)3Sn 9306 (C4H9)3SnCH 0.7a = Molecular weight and all fragment ion assignments are based on49Table 2.3 Major fragment ions for dibutyldimethyltm and tricyclohexylmethyltin.Standard compound M/Z Fragment ion Relative Intensity(%)(C4H9)2Sn(CH3 121 SnW 35(MW = 264) 135 CH3Sn 70150 (CH3)2Sn 99177 C4H9Sn 14193 C4H9SnHCH3 52207 C4H9Sn(CH3)2 100249 (C4H9)2SnCH3 39264 (CSn(CH 5(C6H1,)3SnCH 121 SnW 17(MW = 384) 135 CH3Sn 52219 C6H11SnHCH3 100301 (C6H11)2SnCH3 98369 (C6H11)3Sn 3384 (C6H,,)3SnCH 4a = Molecular weight and all fragment ion assignments are based on ‘20Sn.50Table 2.4 Major fragment ions for dicyclohexyldimethyltin and tetrapropyltin.Standard compound MIZ Fragment ion Relative Intensity(%)(C6H1,)2Sn(CH3) 121 SnH4 13(MW = 316) 135 CH3Sn’ 35150 (CH3)2Sn 94203 C6H11Sn 5219 C6H11SnHCH3 15233 C6H11Sn(CH3)2 100301 (C6H11)2SnCH3 10316 (C6H11)3Sn 19(C3H7)4Sn 121 SnH 46(MW 292) 135 CH3Sn 9163 C3H.Sn 91207 (C3H7)2SnW 91249 (C3H7)Sn 100292 (C3H7)4Sn 3a = Molecular weight and all fragment ion assignments are based on 120Sn512.4.3 GC-MS elution profile and masses of fragment ions used for selected ionmonitoring.The elution profile of the standard tetraorganotin compounds is shown inFigure 2.5. The retention times and the retention time window (the time framewithin which the retention time can vary and still be valid) used for the GC-MSanalyses of the organotin compounds are shown in Table 2.5.Table 2.5 Retention time and retention time window used for (3C-MS SIManalysis.Compound Retention time Retention time window(mm) (mm)(C4H9)2Sn(CH3 12.42 11.50-13.33(C3H7)4Sn 13.97 13.33-14.25(C4H9)3SnCH 14.60 14.25-15.50(C6112Sn(CH 16.13 15.50-17.00(C6H11)3SnCH 18.93 17.00-20.00The fragment ions chosen for GC-MS SIM were selected on the basis of theirhigh intensities in low resolution mass spectrometry (Tables 2.2, 2.3, 2.4). Thefragment ions and masses monitored for each organotin compound are shown inTable 2.6.5211:29 13:10Retention Time (min:sec)16:33 18:14 19:56Figure 2.5 GC-MS elution profile of the standard tetraorganotin compounds.Peaks a, b, c, d, e, correspond to dibutyldimethyltin, teirapropyltin,tributylmethyltin, dicyclohexyldimethyltin, and tricyclohexylmethyltmrespectively.14:511 100 200 300 400 500Scan Number53For each fragment ion, masses corresponding to tin 116, 118 and 120 isotopes weremonitored (Table 2.6).Table 2.6 Fragment ions and masses used to detect and quantify each organotincompound in GC-MS SIM.Compound M/Z monitored Fragment ion(C4H9)3SnCH 193, 191, 189 C4H9SnHCH3(C4H9)2Sn(CH3 207, 205, 203 C4H9Sn(CH3)2(C6H1.,)3SnCH 219, 217, 215 C6H11SnHCH3(C6H,1)2Sn(CH3 233, 231, 229 C6H11Sn(CH3)2(C3H7)4Sn 249, 247, 245 (C3H7)Sn2.4.4 Suitability of tetrapropyltin as internal standard as studied by using gaschromatography (GC).The use of internal standards eliminates the effect of variations in theinstrument’s operating parameters on the analyte. In GC or GC-MS, variations ininjection volume, ion source temperature, carrier gas flow rate, could causeconsiderable errors from run to run, if not eliminated by the method of internalstandardization. Ideally, compounds used as internal standards should be structurally54similar to the analyte and their spectroscopic response should also be similar to thatof the analyte. Also, the internal standard should not be naturally present in theanalyte. Tetrapropyltin satisfies these conditions closely. No natural source oftetrapropyltin in the marine environment is known, and its gas chromatographicretention time is in the range of the retention times of the analytes (Fig 2.5). Thisensures that the internal standard and the analytes experience similar broadeningeffects on the capillary column. Also, calibration graphs obtained for tributylmethyltinsolutions by using different concentrations of tetrapropyltin (2- 50 tg/mL as Sn) asinternal standard, gave high correlation coefficients (Table 2.7).Table 2.7 Regression data for graphs obtained with various concentrations ofthe internal standard tetrapropyltin.Conc. (C3H7)4Sn Regression equation for Regression(/.Lg/mL as Sn) (C4H9)3SnCH coefficient2 Y=0.4964X - 0.0924 0.99804 Y=0.2514X-0.0521 0.99106 Y=0.1659X - 0.0322 1.000050 Y=0.0190X - 0.0023 1.0000The calibration curves obtained for tributylmethyltin by using differentconcentrations of the internal standard are shown in Figures 2.6 and 2.7. The error55aeIc.)0I5.04.54.03.53.02.52.01.51.00.50.00 2 4 6 8 10Conc.(H,)3SnCH (ppm Sn)12(a)(b)0 2 4 6 8 10 12Conc. (CH9)3SnCH (ppm Sn)Effect of internal standard concentration on the linearity of calibrationcurves for tributylmethyltin. Graphs a and b were obtained by using 2jtglmL (as Sn) and 4 jgImL (as Sn) tetrapropyhin respectively.aIFigure 2.656CiDc-)C-.Figure 2.7 Effect of internal standard conceniration on the linearity of calibrationcurves for tributylmethyltin. Graphs a and b were obtained by using 6g/mL (as Sn) and 50 LgImL (as Sn) tetrapropyltin respectively.0 2 4 6 8 10 12Conc. (CH9)3SnCH (ppm Sn)(a)(b)0200.150.100.050.00CIDCL.0 2 4 6 8 10Conc. (CH9)3SnCH (ppm Sn)1257bars in Figures 2.6 and 2.7 represent the standard error for three replicatedeterminations. All the calibration graphs for tributylmethyltin obtained by using thevarious concentrations of tetrapropyltin (2 tg/mL to 50 ig/mL as Sn) as internalstandard gave good linearity (Figures 2.6 and 2.7). Thus, the internal standard couldbe applied in concentrations up to 50 tg/mL (as Sn) without introducing nonlinearity in the calibration curves. The regression equations obtained for the graphsare shown in Table 2.7.An ideal internal standard would have been an analogous stable isotope of thebutyltin or cyclohexyltin compounds because it would be chemically very similar tothe analyte. Deuterated tributylmethyltin was examined for use as internal standardin GC-MS SIM, but was found to be unsuitable because it caused a decrease in theinstrument’s sensitivity to tributylmethyltin detection as explained in Section 2.3.1.3.2.4.5 Detection limit, calibration curves, and precision for the GC-MS SIM analysis.2.4.5.1 Detection limit and calibration curves obtained by GC-MS SIM.The limit of detection was obtained from a plot of the peak area ratio of thestandard tetraorganotin compounds to the internal standard versus the concentrationof the standard tetraorganotin solutions (Figures 2.8 and 2.9). The error barsrepresent the standard error for three replicate determinations.The detection limit defined as the analyte concentration which gives a signalequal to the signal of the blank, plus thrice the standard deviation of the blank, was58__16141210rJD‘‘ 86430Ci20025201510Conc. (C4H9)2Sn(CH3(ppm Sn)0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4Conc. (CH9)3SnCH (ppm Sn)(a)(b)Figure 2.8 GC-MS calibration curves for (a) dibutyldimethyltm and (b)0.0 0.2 0.4 0.6 0.8 1.0 1.2tributylmethyltin.ciciCCl)Nci©CtCtC?I..CtC?CCl)3.5cici 2.5Cl)nI2.0ci— 1.5CCtZ 1.0Ct-C?I... 0.5.40.05910864204.00.0 0.2 0.4 0.6 0.8Cone. (C6H11)5n(CH31.0 1.2 1.4(ppm Sn)3.0(a)(b)0.0 0.2 0.4 0.6 0.8 1.4Cone. (C6H11)35n fJFigure 2.9 GC-MS calibration curves for (a) dicyclohexyldimethyltin and (b)1.0 1.2(ppm Sn)tricyclohexylmethyltin.60calculated according to the method previously described by Miller and Mi1ler1 fromthe calibration curves (Figures 2.8 and 2.9). According to these authors1M, at thelimit of detection the analyte signal is given by the equation Y=Y8 + 3S where Yis the analyte signal at the limit of detection, Y8 is the blank signal and S8 is thestandard deviation of the blank. The standard deviation of the blank SB can becalculated as the standard deviation of the y residuals The blank signal Y8 maybe taken to be the intercept of the graph on the y axis. From the working calibrationcurves of the analytes (Figures 2.8 and 2.9), the intercept of the graph on the y-axisis obtained, and the standard deviation of the y-residuals is calculated. Then, thedetection limit is determined (Table 2.8).Table 2.8 Detection limits for organotin compounds by GC-MS SIM.Compound Detection limit(j.ig/mL as Sn)(C4H9)3SnCH 0.053(C4H9)2Sn(CH3 0.028(C6H11)2Sn(CH3 0.049(C6H11)3SnCH 0.06461Typical calibration curves obtained by the least squares method, for the quantitationof the organotin compounds from environmental samples by GC-MS SIM are shownin Figures 2.8 and 2.9. A linear relationship is obtained over the concentrationTable 2.9 Calibration equations used for the quantitation of environmentalsamples by GC-MS SIM.Compound Calibration equation Regressioncoefficient(C4H9)3SnCH RA = 4.0378RC - 0.1554 0.9944(C4H9)2Sn(CH3 RA = 2.9721RC - 0.5492 0.9989(C6H11)2Sn(CH3 RA = 1.5812RC - 0.1093 0.9817(C6H,1)3SnCH RA = 0.6196RC - 0.1742 0.9685RA =Relative peak area ratio of organotin compound to internal standard,and is plotted as y-axis.RC =Relative concentrations of organotin compound to internal standard, andis the x-axis.range studied. The regression equations obtained from a direct plot of the peak arearatios of the tetraorganotin compounds to the internal standard versus the variousconcentrations of the tetraorganotin compounds were modified to obtain the working62calibration equations by substituting RC (concentration ratio of tetraorganotin tointernal standard) for X in the general form of a straight line equation Y=MX + C.The resulting calibration equations are shown in Table 29.An obvious characteristic of the calibration curves in Figures 2.8 and 2.9 isthe failure of the regression line to pass through the graph’s origin despite variousoptimization steps in the GC-MS operating conditions. This phenomenon was veryreproducible in calibration curves obtained at various times and consequently isunlikely to affect its use for quantitation. A possible consequence of this effect is thedifficulty in attaining a very low detection limit required for trace metal speciation.2.4.5.2 Precision of the GC-MS SIM method.The precision of the GC-MS SIM analysis was determined by analyzingreplicate injections of mixtures of the standard tetraorganotin compounds and theinternal standard dissolved in n-heptane. The precision was determined at twoconcentrations 0.2 g/mL and 1.2 j.ig/ mL (as Sn). The relative standard deviationfor six replicate injections was calculated (Table 2.10). There was no significantdifference in precision at the two organotin concentrations as tested by means of atwo-tailed F-test at 5% probability level163.The sensitivity of the GC-MS SIM method as shown in the slope of thecalibration graphs (Table 2.9) follows the order tributylmethyltin >dibutyldimethyltin > dicyclohexyldimethyltin > tricyclohexylmethyltin.63Table 2.10 Precision for six replicate injections of organotin compoundsas determined by (JC-MS SIM.Compound Precision (RSD %)0.2ig/mL as Sn l.2g/mL as Sn(C4H9)3SnCH 5.4 8.1(C4H9)2Sn(CH3 9.4 10.5(C6H11)2Sn(CH3 11.1 10.0(C6H1,)3SnCH 14.8 7.42.4.6 Recovery studies on the extraction procedure.Recovery studies of the organotin compounds were carried out at theconcentration level of 1.5 tg (as Sn) per 40 g (wet wt) of shrimp as described insection 2.3.4 above. The extraction procedure affords good recoveries for tributyltin,dibutyltin and dicyclohexyltin species (Table 2.11). The detection of thetricyclohexyltin species was hampered by the elution of other unidentified compoundsfrom the sample matrix in its retention window (Fig 2.10). Thus in Figure 2.10, thereis an unidentified peak E, of very high intensity which elutes at the same retentiontime as tricyclohexylmethyltin and completely masks the peak due to this tincompound. Therefore, the quantitation of tricyclohexyltin was not carried out. The64other peaks labelled A, B, C and D on the diagram (Figure 2.10) are due todibutyldimethyltin, tetrapropykin, tributylmethyltin and dicyclohexyldimethyltinrespectively. The unlabelled peaks in Figure 2.10 are unknown and their massspectra do not show tin isotope pattern. The mass spectra for peaks A, C and D(Figure 2.10) are shown in Figures 2.11 and 2.12 (i) and (ii) respectively.Table 2.11 Recovery of organotin compounds spiked into Shrimp (Pandalustridens) by extraction with methylene chloride.Compound Percentage recovery ± (RSD %)(C4H9)3SnCH 96.9 ± 2.1(C4H9)2Sn(CH3 99.9 ± 1.6(C6H11)2Sn(CH3 93.0 ± 9.1a = Relative standard deviation of two extractions, each of tworeplicate injections into the GC-MS.RSD = Relative standard deviation.65Figure 2.10100Retention Time (min:sec)16:33 18:14200 300Scan NumberSelected ion current chromatogram of standard organotm compoundsspiked into shrimp. Peak B corresponds to the internal standard.2070100Figure 2.11150 200 250 300 350 400MJZMass spectra of peak A in Figure 2.10. The peak at m/z207 indicatesthe presence of dibutyltin.66100 - 193(i)50-zz—I I I l I I I I I I I I I I I100 150 200 250 300 350 400MJZ100(ii)50z0—111111111 III 1111111 lilt 111111111 11111111100 150 200 250 300 350 400MuFigure 2.12 Mass spectra of (i) peak C and (ii) peak D in Figure 2.10 aboveindicating the presence of tributyltin and dicyclohexyltin speciesrespectively.672.4.7 Organotin concentrations in some marine organisms of British Columbia,CadaThe marine animals selected for organotin analyses consisted mainly ofbivalves molluscs. This was mainly because of their availability in the marinelocations sampled. The analysis of marine organisms for organotin compounds wascarried out with the following objectives:(i) To obtain an indication of the geographical spread of organotin pollution inthe coastal areas of British Columbia in terms of dibutyltin, tributyltin, and thecyclohexyltin compounds. The latter were of particular interest becausedicyclohexyltin species have been found in surface microlayers of some BritishColumbian marine waters, and biota117.(ii) To study organotin concentrations in particular species of marine animals,over a period of three years.(iii) To examine organotin concentrations in different animals from one particularlocation to see if any particular animal has the ability to accumulate organotincompounds more than the others.In practice, animal availability varied from location to location thereby makingit difficult to obtain data from the same species of animals in all locations sampled.The map of the locations sampled is shown in Appendix A.2.4.7.1 Organotin concentrations in oysters.The organotin concentrations in the Pacific oyster Crassostrea gigas from68some locations in British Columbia, Canada are shown in Table 2.12.Table 2.12 Organotin concentrations in the oyster Crassostrea gigas fromsome coastal areas of British Columbia.ng/gasSn(Wet weight)Year of Location (C4H9)2Sn (C4H9)3Sn (C6H1,)2SncollectionJuly, 1991 Denman 12.5 ± 1.8 ND NDIslandbSept., 1991 Von Donop 17.3 ± 0.3 7.6 ± 0.3 NDInlet, CortesIslandMay, 1991 Pendrell ND ND NDSoundND = Not detected.a = Standard error for two separate sample determinations. All otherstandard errors given are for two replicate injections of one sample.b= Oysters were purchased.I69Oysters in particular have been shown to be very sensitive to tributyltinpollution24.Effects of tributyltin pollution on oysters include shell malformation andretarded growth22. Oysters from Pendrell Sound British Columbia (Table 2.12)showed no organotin pollution. Comparative data on the organotin body burden• ofoysters from another Canadian location, Fanny Bay, British Columbia has beenreported by Stewart and Thompson1M.According to them, the oyster Crassostreagigas contained tributyltin and monobutyltin concentrations of 52 ng/g (dry wt as Sn)and 4.6 ng/g (dry wt as Sn) respectively. These organotin concentrations translate toapproximately 10.4 ng/g (wet wt as Sn) tributyltin species and 0.92 ng/g (wet wt asSn) monobutyltin species. No dibutyltin species were detected by the authors”. Thisshows that Fanny bay is comparable to Von Donop Inlet, British Columbia intributyltin pollution (Table 2.12). Other organotin data for oysters from other partsof the world are available. Rice Li165 have reported tributyltin concentrations of9 ng/g (wet wt as tributyltin cation) or 3.7 ng/g (wet wt as Sn) for oysters from SarahCreek, and 834 ng/g (wet wt as tributyltin cation) or 341.3 ng/g (wet wt as Sn) foroysters from Kings Creek, Virginia U.S.A.. Tributyltin concentrations of 49.74 - 189ng/g (wet wt as tributyltin cation) or 20.4 - 77.3 ng/g (wet wt as Sn) have also beenreported by Wolniakowski et al1 for the oyster Crassostrea gigas from Coos BayEstuary, U.S.A..A comparison of the butyltin body burden for oysters analyzed in this study(Table 2.12) with the butyltin body burden reported by Rice et al165 indicates thatoysters from Von Donop Inlet, British Columbia, contain higher tributyltin levels70than those from Sarah Creek, Virginia U.S.A., but less tributyltin levels than oystersfrom Kings Creek, Virginia, U.S.A..Concentrations expressed in wet weight are not comparable to concentrationsexpressed in dry weight, unless appropriate conversion factors are applied. Anapproximate conversion factor applied in this study for oysters is x p.glg dry wt basis= 5x .tg/g wet wt basis. This conversion factor was arrived at, after freeze-dryingknown weight of wet oyster samples.Waldock and Miller167 have reported tributyltin levels of up to 4.5 ig/g dryweight or approximately (0.37 jg/g wet wt as Sn) for some oysters from England.Rapsomanikis and Harrison have also reported tributyltin levels of 0.027 - 1.66jig/g (dry wt as tributyltin cation) or 2.2 - 135.9 ng/g (wet wt as Sn) and dibutyltinlevels of 0.012 - 0.402 tg/g (dry weight as dibutyltin cation) or 2.4 - 32.9 ng/g (wetwt as Sn) for some oysters from England. Other tributyltin levels in oysters have beenreported by Stewart and de Mora for the Mangrove oyster Crassostrea mordax ofFiji. Tributyltin concentrations in the range 626 to 3180 ng/g (dry wt as tributyltincation) or 51.2-260.3ng/g (wet wt as Sn) were obtained. In New Zealand, tributyltinconcentrations in the range 0.033 - 1.38 g/g (dry wt as tributyltin) or 2.7- 110.5ng/g (wet wt as Sn) and 0.049 - 0.467 tg/g (dry wt as tributyltin cation) or 4.0- 38.2ng/g (wet wt as Sn) have been reported for the oysters Crassostrea gigas andSaccostrea glomerata respectively, for the Tamaki Estuary of New Zealand169. Hanand Weber79 have determined dibutyltin and tributyltin concentrations of 840 and2200 ng/g (dry wt as Sn) respectively for a French oyster sample. By using a dry71weight/wet weight conversion ratio of about 0.2 as given by these authors7’9,dibutyltin and tributyltin concentrations of 168 and 440 ng/g (wet wt as Sn)respectively were obtained for the French oyster sample.A visual inspection of the oysters’ shells prior to their analysis did not revealobvious shell malformations. Oysters from the Canadian location Denman Island didnot show the presence of tributyltin species, but did show the presence of dibutyltinspecies which may have originated from the metabolism of tributyltin compounds.The available data in Table 2.12 do not indicate pollution by dicyclohexyltin species.2.4.8 Spread of organotin compounds in the Canadian environment.The extent of organotin pollution in British Columbia was monitored bysampling various available marine organisms from different locations in BritishColumbia. The results obtained are shown in Table 2.13 below. The concentrationsand species of organotin compounds detected in any one location should be areflection of the type of industrial activity going on in that environment. By the verynature of introduction of tributyltin compounds into the environment, areas of highboating or shipping activity should show high tributyltin concentrations. Data in Table2.13 show the occurrence of tributyltin species in a substantial number of locationssampled. An interesting feature of Table 2.13 is the observation that the highestconcentration of dibutyltin species 67.3 and 39.6 ng/g (wet wt as Sn) were found inthe Blue mussels Mvtilus edulis from Anyox shore, and Kitimat respectively. Thehighest concentration of tributyltin species (37.3 ng/g wet wt as Sn) was also found72Table 2.13 Organotin concentrations, spread and speciation in some marinelocations of British Columbia.Organism Location and Conc.(nglgdate of wet wt Sn)collection (C4H9)2Sn (C4H9)3Sn (C6H11)2SnBlue mussel Marklane 39.6 ± 0.6 ND(Mytilus edulis) Point, Kitimat(1990)Soft shell Clam(Mya arenaria)Dundas Island ND(1990)19.4 ± 0.7 NDShrimp(Pandalus tridens)Holberg NDSound(1991)Blue mussel(Mytilus edulis)Wreck Beach, 6.7 ± 0.1Vancouver(1989)14.4 ± 0.4 3.5 ± 0.1Bentnose Clam(Macoma nasuta)Table 2.13 continuedAlice Arm ND(1989)on next page.NDND NDND ND73Table 2.13 continued.Organism Location (C4H9)2Sn (C4H9)3Sn (C6H1,)2Sn& date of (nglg wet wt (ng/g wet wt (ng/g wet wtcollection as Sn) as Sn) as Sn)Bentnose Clam Hilton ND ND ND(Macoma nasuta) Point,Kitimat.(1990)Soft shell Clam Hastings ND ND ND(Mya arenaria) Arm(1990)Blue mussel Anyox 67.3 ± 8.6 37.3 ± 15.0 21.3 ± 0.1(Mytilus edulis) Shore(1990)Soft shell Clam Anyox 1.9 ± 0.2 0.7 ± 0.2 ND(Mya arenaria) Shore(1989)Basket Cockles Anyox ND 7.7 ± 2.8 ND(Clinocardium Slag shorenuttallii) (1989)74Table 2.13 contd.California mussel Quatsino ND 9.2 ± 1.5 ND(Mytilus Soundcalifornianus) (1990)ND = Not detectedin the same Blue mussels from Anyox shore. Blue mussels from Wreck beach,Vancouver, also have moderately high concentrations of tributyltin species (14.4 ng/gwet wt as Sn) when compared to other organisms studied, except the Soft shell clamMya arenaria from Dundas Island. There appeared to be a tendency for Blue musselsto accumulate relatively higher concentrations of organotin compounds than theother bivalve molluscs studied. The occurrence of dicyclohexyltin species is notwidespread. Dicyclohexyltin species were found only in the Blue mussels from WreckBeach, Vancouver, (Figure 2.13) and Anyox shore. Also interesting, is the absenceof dicyclohexyltin species in clams and cockles from the same Anyox location. Itseems possible that Blue mussels have greater ability to accumulate dicyclohexyltincompounds than the other bivalves studied. This may indicate the incapability of Bluemussels to effectively metabolize or excrete dicyclohexyltin compounds, therebysuggesting different metabolic pathways between blue mussels and other bivalves withregard to organotin metabolism. If this relation holds true, mussels may become goodbiological indicators for monitoring cyclohexyltin pollution. An examination of Fig752.13(a), demonstrates the superiority of mass spectrometric detection over most nonspecific detectors. With non-specific detectors, the dicylohexyltin peak D, could havebeen discarded as baseline noise.Apart from the present work, and an earlier report by Cullen et al1’7, noconcentrations of dicyclohexyltin species have been reported for mussels in BritishColumbia. However, tricyclohexyltin concentration of 36 ng/g (dry wt as Sn) hasbeen found for sediments from Esquimalt Harbour, British Columbia170. Recently,cyclohexyltin species have been reported for environmental samples from St John’sharbour, New Foundland, Canada’7’ and Spain172. Other butyltin concentrations inthe range obtained in this study have been found by Garrett’73 (Table 2.14) for Bluemussels from Nanoose Bay and Wood Bay, British Columbia, Canada. Stallard Li122 have reported dibutyltin concentrations in the range 0.087 - 0.169 .ig/g (wet wtas dibutyltin cation) and tributyltin concentrations in the range 0.068 - 1.067 j.ig/g(wet wt as tributyltin cation) for Blue mussels from San Diego bay, U.S.A. (Table2.14).Higashiyama et al174 have also reported dibutyltin and tributyltinconcentrations in the range 0.04-0.54.ig/g, and 0.02-0.24g/g (wet wt as organotincation) respectively for mussels from Tokyo bay, Japan (Table 2.14).To afford a comparison of the British Columbian mussels with the reportedorganotin concentrations in Table 2.14, the organotin concentrations for mussels inthis study were converted from ng/g (wet wt as Sn), to g/g (wet wt as organotincation) in Table 2.15. A comparison of organotin data in Tables 2.14 and 2.1576Retention Time (min:sec)11:29 13:09 14:51 16:32 18:14 19:55100 I I I(a)Cl)Z501 100 200 300 400 500Scan Number100 233(b)F010 I100 150 200 250 3Ô0 350 400MIZFigure 2.13 (a) Selected ion current chromatogram of extract from Blue musselMytilus edulis from Wreck Beach, Vancouver. (b) Mass spectra of peakD, revealing the presence of dicyclohexyltin species.77Table 2.14 Some organotin concentrations reported for the Blue mussels Mytilusedulis..tg/g (wet wt asorganotincation)Location (C4H9)2Sn’ (C4H9)3Sn (C4H9)Sn3 ReferenceU.S.ASan Diego Bay 0.169-0.087 0.068- 1.067 0.076-0.257 Stallarda1122JapanTokyo Bay 0.04- 0.54 0.02 - 0.24 0.02- 0.12 Higashiyamaet al17’4CanadaNanoose Bay 0.002 0.007 0.003 Garret17’3Wood Bay 0.02 0.037 0.003 Garret173indicates that the concentration of tributyltin species found in Anyox, BritishColumbia is in the range reported by both Stallard j122 and Higashiyama Li’74for San Diego Bay, U.S.A and Tokyo Bay, Japan respectively. The level of tributyltinspecies present in Anyox British Columbia is higher than the levels in both NanooseBay, and Wood Bay British Columbia respectively. The occurence of dibutyltin78species in Blue mussels from Markiane point, Kitimat and Wreck Beach, Vancouverwithout a corresponding occurence of the tributyltin species (Table 2.15) issurprising, and may indicate that for Blue mussels, the excretion of dibutyltin speciesTable 2.15 Organotm concentrations in the Blue mussel Mytilus edulis convertedto pgIg wet wt as organotin cation.Organism pg/g Wet wt. (asand (Location) Organotin cation)(C4H9)2Sn4 (C4H9)3Sn (C6H11)2SnBlue mussel 0.08 ND ND(Markiane Point,Kitimat)Blue mussel 0.01 0.04 0.01(Wreck beach,Vancouver)Blue mussel 0.13 0.09 0.05(Anyox shore)is slower than that of tributyltin species. For soft shell clams, the opposite trend wasobserved because dibutyltin species were generally not detected (Section 2.4.10,Table 2.18).792.4.9 Organotm concentrations in various organisms from the same locations.Marine animals from the same locations were sampled for the presence oforganotin compounds with a view to finding their distribution among organisms. TheTable 2.16 Organotm distribution in marine animals from Camano Sound,British Columbia.Organism Year of Conc. (ng/g ascollection Sn wet wt)(C4H9)2Sn (C4H9)3Sn (C6H11)2SnButter clam 1989 ND ND ND(Saxidomusgiganteus)Basket cockle 1989 ND ND ND(Clinocardiumnuttallii)Soft shell clam 1989 ND 7.0 ± 2.6a ND(Mya arenaria)a= Standard error for two determinations, of two replicate injections each.ND = Not detected.80organotin distribution in organisms from Camano Sound and Tasu Sound is shownin Tables 2.16 and 2.17 respectively. In Camano Sound, no dibutyltin ordicyclohexyltin compounds were detected in the three animals sampled. Only the Softshell clam Mya arenaria showed the presence of tributyltin species (Table 2.16). Theoccurrence of tributyltin species in the Soft shell clam, without a correspondingoccurrence in the butter clam is surprising, and may indicate different mechanismsof tributyltin detoxification even in different species of the same animal. Of the twoanimals from Tasu Sound (Table 2.17), the little neck clam showed a much higherconcentration of tributyltin species than the Blue mussel. Unfortunately, little neckclams from other locations were not available to study this trend further.Table 2.17 Organotin distribution in marine anima1 from Tasu Sound, BritishColumbia.Conc. ng/g asSn (wet wt.)(C4H9)2Sn’ (C4H9)3Sn’ (C6H11)2SnBlue mussel ND 13.2 ± 3.1 ND(Mya arenaria)Native littleneck clam ND 177.0 ± 11.9 ND(Protothaca staminea)812.4.10 Distribution of organotin compounds in marine anim2ls studied overa period of three years.The concentration and speciation of organotin compounds in Soft shell clamsfrom Quatsino Sound, British Columbia were monitored for over a period of threeyears (Table 2.18). An examination of the data in Table 2.18 indicates a veryTable 218 Organotin body burden for Soft shell clams Mya arenaria fromQuatsino Sound, British Columbia studied over a period of threeyears.Year of collection ng/g Sn (wet wt)()a(C4H9)2Sn (C4H9)3Sn (C6H11)2Sn1989 ND 26.3 ± 0.6 ND1990 ND 12.8 ± 0.4 ND1991 ND •s ± 18b NDa=Standard error for two replicate injections.b=Standard error for two separate determinationssignificant decrease in the concentrations of tributyltin species with time. Such a verysignificant decrease in tributyltin concentration could only be possible if the inputsource of tributyltin compounds in this location is decreasing with time. According82to Maguire155, in 1989 the Canadian Government regulated tributyltin compoundsunder the Pest Control Products Act (Canada Department of Agriculture 1989).Under this regulation, the permitted daily release rate of tributyltin species is 4jtg persquare centimetre of hull surface. This regulation also prohibits the use of antifoulingpaints containing tributyltin compounds on vessels less than 25 metres in length. Assurmised by Maguire155,these regulations should minimize the environmental impactof antifouling uses of tributyltin compounds in Canada. The decreasing concentrationof tributyltin species with time as shown in Table 2.18 may represent the impact ofthe Government’s regulation on the input of tributyltin compounds into the marineenvironment. An important trend that is observable from Table 2.18 is the absenceof dibutyltin species in the clams. Usually dibutyltin species should co-exist withtributyltin species in organisms because dibutyltin species are metabolites oftributyltin compounds in animals. This trend may indicate that Soft shell clamsgenerally do not metabolize tributyltin species or that dibutyltin species are veryquickly excreted from the clams. This observation is in contrast to the trend foundfor blue mussels (Table 2.15), where all the Blue mussels analyzed containeddibutyltin species. This observation pQints to different detoxification mechanisms forsoft shell clams and blue mussels.The occurrence of organotin species in some remote British Columbianlocations such as Anyox, Hastings Arm, Alice Arm, and Tasu Sound is surprisingbecause of the very low boating and agricultural activity in these locations: perhaps,aerial transport of these compounds needs to be considered.83CHAPTER 3EFFECT OF TRIBUTYLTIN CHLORIDE, MONOBUTYLTINTRICHLORIDE AND TRIM[THYLTIN HYDROX[DE ON THEPERMEABILiTY OF EGG PHOSPHATIDYLCHOLINE LIPOSOMES.3.1 INTRODUCTIONThis chapter describes the effect of some organotin compounds namelytributyltin chloride, monobutyltin trichioride, and trimethyltin hydroxide on modelbiological membranes formed by the hydration of egg phosphatidyicholine (EPC) ora mixture of organotin compound and EPC in tris buffer to form liposomes, alsoknown as vesicles. The experiment was originally designed to study the permeationof these organotin compounds through these liposomes. However at the highconcentrations of tributyltin chloride and monobutyltin trichioride needed inside theliposomes for their 1H NMR signals to be observed, the liposomes do not form.Therefore, the approach adopted was to use a molecular probe which is capable ofeasy permeation through the liposomes, to monitor the effects of low concentrationsof organotin compounds on the permeability of these model biological membranes.The compound chosen as a probe was dimethylarsinic acid (DMA). The reasons forchoosing DMA as a probe molecule are given below in Section 3.2. In an earlierstudy by Tosteson and Weith156 the probe ions tetraphenylboron, andtetraphenylarsonium were used to study the effect of tributyltin chloride on themembrane potential of a phosphatidylethanolamine planar lipid bilayer. A decrease84of about 70 mV in the intrinsic dipole potential of the planar lipid bilayer caused bytributyltin chloride was observed by Tosteson and Weith156.The experimental technique employed in the present study was NMRspectroscopy. NMR spectroscopy is well suited for the study of solute or molecularpermeation through liposomal membranes, provided the NMR signals of the soluteinside and outside the liposomes can be differentiated from each other. Thedifferentiation of the outside and inside NMR signals is usually achieved by the useof spectroscopic shift reagents; these reagents are usually first row transition metalcomplexes and complexes of the lanthanide elements1, and are usually added to thesample prior to the NMR experiment. By using NMR spectroscopy the permeationof molecules or solutes can be followed to equilibrium without the intermittentwithdrawal of samples from the reaction system. In addition, the technique is capableof providing information on the state of the liposomes, particularly liposome lysisduring the experiment, because the NMR signal inside and outside the liposomewould collapse into a single peak if the liposome disintegrates or bursts.3.2 DIMETHYLARSINIC ACID (DMA) AS A PROBE FOR STUDYING THEEFFECT OF ORGANOTIN COMPOUNDS ON THE MEMBRANES OFLIPOSOMES.The permeation of dimethylarsinic acid (DMA) through EPC liposomes hasbeen studied by Herring ci176. DMA has the following properties which make itsuitable as a probe molecule for further studies:85(i) DMA permeates across EPC liposomes by passive diffusion(ii) DMA has good aqueous solubility which enables high concentrations to betrapped in the small aqueous volumes of the liposomes. This in turn makes it easyto observe the NMR signals of DMA in the liposomes. Butyltin compounds do notpossess high enough aqueous solubility to enable the NMR signals of trapped butyltincompounds to be detected.(iii) The rate of efflux of DMA from EPC liposomes is slow enough to permit itsstudy by NMR spectrometry.(iv) The methyl hydrogen atoms of DMA give rise to a simple NMR spectra(singlet) which can be shifted by using spectroscopic shift reagents.3.3 LIPOSOMES AS MODELS FOR BIOLOGICAL MEMBRANES.Biological membranes are made up of two major components, phospholipidsand proteins”’’, and other components such as oligosacchrides178.The major barrierto membrane permeability is provided by the phospholipid bilayer. The proteins areinserted into the phospholipids which are oriented in the bilayer. An importantproperty of the phospholipids is the possession of hydrophillic and hydrophobic ends(FiHxre 3.1). WhHx hydrated in aqueous solutions, most phospholipids form closedstructures called vesicles or liposomes which possess internal aqueous volumes(intraliposomal compartment) which can be used to trap many compounds (Figure3.2). The permeability properties of the liposomes are similar to those of biologicalmembranes”79. The advantage of using liposomes over biological membranes for86permeability studies is that experimental results are easier to interpret becauseproteins and oligosacchrides, which might otherwise complicate permeation processesN(CH3)(a) CH2CH2ci0H CH2H—C C—H0=o c=oCH CH, (b)CH, H.-CH2CN3 EthanolamineCH2 CH2: : -CHCH(NH) OOH H -CH2CH(OH)CHO GlycerolCH,, C1-1.Figure 3.1 (a) Structure of a phospholipid (phosphatidyicholine) and (b) othercommonly occuring head groups on the phospholipid.87are absent. Also, the liposomes can easily be prepared in a controlled andreproducible manner.3.4 TYPES OF LIPOSOMES AND METHODS OF PREPARATION.Various methods for the preparation of liposomes are available. Thesemethods of preparation have been a subject of reviews by Hope LJ180,and SzokaQcExtraliposomalc5 D volumeFigure 3.2 Liposome, showing the intraliposomal compartmentlvolume wheremolecules of a permeant can be encapsulated.and Papahadjopolous181. Three types of liposomes namely multilamellar vesicles(MLVs), small unilamellar vesicles (SUVs), and large unilamellar vesicles (LUVs)have been popularly used and the methods available for the preparation of these88vesicles are given below.3.4.1 Multilamellar vesicles (MLVs).Bangham prepared the first vesicles (MLVs)182 in 1965. His method involveda gentle dispersion of a lipid in buffer. The vesicles that formed were heterogenousin size, and aqueous volumes of the different lamellae were later reported by Grunerto be depleted in solutes relative to the buffer in which they are made, andas such, are under osmotic compression. MLVs having uniform solute distribution inthe lamellae can be prepared by the methods reported by Gruner j183 and Mayeret a1. The methods involve the evaporation of ether from an ether-buffer-lipidmixture, followed by resuspension of the sonicated emulsion in buffer183 or therepeated freezing and thawing of the lipid-buffer preparation. Another method ofMLV formation reported by Kirby and Gregoriadis’85, involved the dehydration oflipids from an aqueous solution by using either freeze-drying or direct vacuumevaporation, followed by controlled rehydration. The major draw back in the use ofMLVs for permeation studies is the presence of multilamellae, and the sizeinhomogeneity of the liposomes.3.4.2 Small unilamellar vesicles (SUVs).Early methods employed for the preparation of SUVs were based on thesonication of multilamellar vesicles. According to Johnson1, the size of the SUVsis dependent on the lipid composition, with the vesicle diameter varying from 204 A89for egg phosphatidyicholine (EPC) vesicles to 362 A for EPC vesicles containing 50%cholesterol. Preparation of SUVs can also be accomplished by the French pressmethod of Barenholtz187.The very small trapped volumes of the SUVs (<O.2iLpertmo1 phospholipid) and vesicle instability are the major draw back to their use inpermeation studies.3.4.3 Large unilamellar vesicles (LUVs).Large unilamellar vesicles (LUVs) are the liposomes of choice for mostpermeation experiments, and were the liposomes used in the present study becauseof their unilamellarity and large trapped volumes. The LUVs can be prepared by theethanol injection method of Kremer the reverse phase evaporation methodof Szoka and Papahadjopoulos’89, the ether injection method of Deamer andBangham’90, the detergent dialysis method of Madden191, and the rapid extrusionmethod of Olson 192 Hope LV93 and MeyerThe ethanol injection, ether injection, and the reverse phase evaporationmethods involve the dispersion of lipids in an appropriate organic solvent, and thesubsequent injection into a buffer. The organic phase is evaporated off at the timeof hydration for the ether injection method, removed under reduced pressure for thereverse phase evaporation method, or is diluted into the buffer for the ethanolinjection method. An additional step involving gel permeation chromatography isusually employed to remove organic solvents. The detergent dialysis method involvesthe detergent induced solubilization of the lipid into micelles and the subsequent90removal of the detergent by dialysis. These methods described above are tedious, andusually entrap residual organic solvents or detergents. The presence of residualsolvents in the LUVs is not desirable because it may change the properties of theliposomes.The entrapment of residual organic solvents can be avoided by rapidlyextruding MLVs under low pressure, as reported by Olson et a1192. These authorsalso reported that reverse phase vesicles exhibit greater size homogeneity after lowpressure extrusion through a polycarbonate filter. The use of moderate pressureextrusion to produce defined pore sized, unilamellar vesicles from multilamellarvesicles has been reported by Hope Lili193 and Meyer3.5 TRANSPORT PROCESSES IN MEMBRANES.Solute or ionic transport across biological membranes can be described interms of the following:(i) Simple or passive diffusion(ii) Facilitated diffusion(iii) Active transportA brief description of these transport processes is given in the following sections:3.5.1 Simple or passive diffusion.Passive diffusion occurs when a concentration gradient exists across amembrane. The movement of molecules through the membrane is due to thermal91molecular motion194. The direction of transport is determined by the concentrationgradient, and diffusion is in the direction of lower solute concentration, untilconcentration on each side of the membrane is equalized. Passive diffusion obeysFick’s first law:J--DdXwhere J, is the flux (mol/cm2/s), D, is the diffusion coefficient (cm2/s) , dX is themembrane thickness, and dC/dX is the concentration gradient.In general, the rate of diffusion is determined by the concentration differenceacross the membrane, the molecular size of the permeant, the viscosity and width ofthe membrane, and on temperature. In passive diffusion, it is assumed that lipophilicsolutes penetrate the membrane by dissolving in the hydrophobic layer and thendiffusing across the bilayer, while hydrophilic solutes pass through aqueous pores onthe membrane. This assumption is based on the observation that the rate ofpermeation is non-saturable, and permeation is not inhibited competitively byanalogous compounds of the permeant195. A detailed description of passive diffusionhas been given by Heinz195.3.5.2 Facilitated diffusion.In facilitated diffusion, the transport of the permeant is aided by the presenceof another molecule capable of acting as a carrier or capable of forming channels in92the membrane. The direction of transport is along the concentration gradient andFick’s first law is not obeyed196. The mechanism of solute transport by facilitateddiffusion is described in terms of the following models:3.5.2.1 Solute iranslocation through channels.In this model, the permeant moves across the membrane via channels.Channels are transient pores formed in the membrane by ionophoric substances. Thetransient pores appear to oscillate between two conformational states. Channels showspecificity, for different permeants, and the specificity shown is not related to the sizeof the permeant. Channels are subject to competitive inhibition.3.5.2.2 Translocation through carriers.The carrier model postulates that a carrier molecule binds specifically to thepermeant molecule at one side of the membrane barrier, transports it through thebarrier, releasing it at the other side. The carrier molecule is able to move freelywithin the bilayer without leaving it. In situations where the size of the carrierexceeds the thickness of the lipid bilayer (30-50 A), it has been suggested that thewhole carrier molecule does not move but only a loose chain or part of the carrierswings from one side of the membrane to the other, releasing the bound permeant’95.A detailed description of facilitated diffusion has been presented by Hofer’94,Heinz195, and Stein197. A schematic diagram of the various steps involved infacilitated diffusion is shown in Figure 3.3.93Lipid bilayer-“-B ABA is the permeantB is the carrier moleculeAB is the carrier-permeant complexFigure 3.3 Schematic diagram of facilitated diffusion (efflux) mediated by acarrier.3.5.3 Active transport.In active transport, a permeant is moved across the membrane by a carriermolecule usually a protein against the permeant’s electrochemical potential gradient.The energy required for this process is provided by ATP hydrolysis, or electron flowconnected with some redox reactions in the cell196. A detailed description of activetransport has been given by Hofer194, and Stein197.943.6 SOLUTE TRANSPORT ACROSS LIPOSOMAL MEMBRANES.3.6.1 Transport of solutes across membranes.When there is a difference in the electrochemical potential of a solute on bothsides of a membrane, there will be a net diffusion of molecules of that solute acrossthe membrane. This situation is represented as follows:-- 0where p. the chemical potential of the solute is given by the following,*+RT a + ZFeV + Vpwhere a, is the activity of the solute, E is the charge on the electron, p is the appliedpressure, V is the volume of the solute, p. is the chemical potential of the solute inits standard state, ‘F is the electric potential, and F is the Faraday’s constant. If theapplied pressure and the electric potential are equivalent on both sides of themembrane, any observed chemical potential difference across the membrane is dueto unequal activity of the solutes on either side of the membrane.The net flux of a solute across a membrane has been described by Stein197,and is given below:ACJ- —D— - -D—8X AXwhere J (mol/s/cm2) is the flux, D (cm2/s) is the diffusion coefficient in the95membrane, and AX is the membrane thickness.The ease of permeation through a membrane is described in terms ofpermeability coefficients. For non-ionic solutes, there is a strong correlation betweenthe permeability coefficieHx for the transport of the solute across a lipid bilayer andthe hydrophobicity of the solute198. This observation is known as Overton’s rule.A schematic diagram for the permeation of a non-ionic solute across aliposomal lipid membrane from the intraliposomal compartment to theextraliposomal compartment is shown in Figure 3.4. As the solute permeates, it alsopartitions between the lipid bilayer and the aqueous phase. An equation that relatesthe permeability of a solute to its partition coefficient has been derived by Jain,and is given below;AXwhere K is the partition coefficient of the solute in the bilayer (Figure 3.4), P (cm/s)is the permeability coefficient, AX (cm) is the membrane thickness and D (cm2/s)is the diffusion coefficient of the molecule in the membrane. The kinetics of thispermeation process is described by the following rate constants:(i) The rate constant for diffusion to the lipid bilayer k1.(ii) The rate constant for diffusion in the lipid bilayer k”.(iii) The rate constant for diffusion away from the lipid bilayer k2.The rate of diffusion in the bilayer is the slowest step and is therefore therate determining step.96Extraliposomal compartmentk2(a)(b)rout — r1 = AXI/Figure 3.4 Passive diffusion of a permeant HA across a liposomal membraneshowing:- (a) the various permeation rate constants and (b) partitioning of thepermeant as it diffuses across.97The permeation of non ionic solutes is influenced by such factors as molecularsize200’1, and hydrogen bonding capability202. Orbach and Finkelstein203 havesuggested that the effect of molecular size and hydrogen bonding capability is lessimportant than the hydrophobicity of the molecule.3.6.2 Transport of ions.In general, the flux of ions across lipid membranes is much lower than the flux ofnon ionic molecules. The permeability of water and ionic solutes across lipidmembranes has been reviewed by Deamer and Bramhall’79.Attempts to explain the low permeability of ions to lipid membranes werecomplicated by the observation that anions permeate lipid bilayers easier thancations. According to Hauser Li204,at pH 5.5 the first order rate constant for theescape of chloride ions from small unilamellar vesicles was three orders of magnitudehigher than the rate constant for sodium ions. Many theories have been put forwardto explain this difference in cation and anion permeabilities. According toPersegian205, the major energetic barrier to membrane transport of ionic solutes isthe Born energy: defined as the energy of an ion in an environment with a givendielectric constant205.Flewelling and Hubbel206 have proposed a mechanism to account for thepermeability difference across lipid bilayers observed for cations and anions.According to them, the observed permeability differences could be accounted for ifother contributions to the Born energy such as image energy, dipole energy, and98neutral energy are considered. Their energy model produced reasonable agreementbetween the observed and calculated thermodynamic parameters for the translocationof tetraphenyiphosphonium cation and tetraphenylboron anion in a lipid bilayer. Theimage energy arises from the interaction between the charge of an ion in the lipidbilayer and the interfaces. The dipole energy arises from a two dimensional array ofpoint dipole sources located at each membrane surface. This dipole source isbelieved to originate from the ester linkages of the fatty acid chain of the lipidbilayer. These dipoles give the interior of the bilayer a positive potential of severalhundred millivolts. This has the effect of increasing the permeability of anions, anddecreasing the permeability of cations. The neutral energy takes into account thenon-electrical interactions between a permeant and the membranes. Such non-electrical interactions include hydrophobicity, and steric factors. However, the Bornenergy considerations are not able to explain the anomalously high permeability ofprotons and hydroxyl ions when compared to other small monovalent ions.On the molecular mechanism of solute and ion transport, no generalagreement has been reached. No one model has satisfactorily explained thepermeation of ionic solutes. For lipophilic solutes, permeation is explained in termsof the solubility-diffusion model, whereby the solute is thought to dissolve in the nonpolar region of the bilayer and cross the bilayer by simple diffusion. For hydrophilicsolutes, permeation is usually explained in terms of diffusion through aqueous poresin the bilayer, or by permeation directly through the lipid bilayer via transient defectswhich occur in the bilayer as a result of thermal fluctuations.993.7 PROPERTIES OF LIPOSOMES CAPABLE OF YIELDINGINVESTIGATWE INFORMATION.The properties of liposomes that can be used to study the effect of othercompounds on the lipid bilayer are its material properties such as permeability,partition coefficient, electrical properties, elastic properties and gel to liquid (L-L)transitions. The material properties of the lipid bilayer have been described byGruner207. These material properties are a function of the vesicle composition, andas such, any interaction of a “foreign” compound such as an organotin compound withthe vesicle would exert some effects on these properties. This has been found to betrue experimentally. Tosteson and Weith’56 have determined that tributyltin chlorideaffects the internal potential of a phosphatidylethanolamine planar lipid membrane,lowering its dipolar potential by 70 mV.In the present study, bilayer permeability was the material property of choiceto be used for studying the effect of tributyltin chloride, monobutyltin trichioride andtrimethyltin hydroxide on the permeability of egg phosphatidyicholine liposomes byusing dimethylarsinic acid (DMA) as a permeability probe.3.8 BUTYLTIN COMPOUNDS: THE NEED FOR THE PRESENT STUDY.The pioneering work of Selwyn et al15’208 on the effect of the triorganotincompounds on membranes has shown that the organotin compounds (trimethyltin,triethyltin, tripropyltin, tributyltin and triphenyltin species) partition into themembranes of mitochondria, liposomes, erythrocytes, and chloroplasts, and mediate100chloride-hydroxide transport across the membrane. Motais et a117 found thattripropyltin chloride is able to act as carrier and mediate chloride-chloride andchloride-hydroxide exchanges in red blood cells.Although the ability of the triorganotin cations to act as carriers for CF andOW in the membrane has been established, the effect of the organotin compoundson the other properties of the membrane has received very little attention. Heywood209 have provided evidence that tributyltin cation associates with the phosphatehead groups at the surface of liposomes. Such interactions are expected to modify themembrane properties of the liposomes. Alteration of membrane properties bytributyltin compounds is suspected to be responsible for the in vitro inhibition ofintracellular Ca2 mobilization reported by Arakawa cAi157.Tosteson and Weith16 have reported that the tributyltin cation preferentiallytransports CF over N03 across a planar lipid bilayer. In the environment, thetoxicity of other pollutants may be enhanced, if tributyltin cation preferentiallymediates their diffusion across biomembranes.Hence, the present study aims to investigate the effect of organotincompounds on the permeability of biomembranes, and any possible transportmediating ability of the organotin cation on a probe permeant; dimethylarsinic acidwhich is also an environmentally occuring compound.1013.9 ThEORETICAL DESCRWrION OF THE DIFFUSION EXPERIMENTAPPLICABLE TO NMR SPECTROMETRY.3.9.1 Passive diffusionDuring the efflux experiments, as the DMA diffuses out of the vesicles, theintensity of the DMA peak inside the liposomes decreases, while the intensity of theDMA peak outside the liposomes increases. A mathematical description for the firstorder efflux of DMA from egg phosphatidyicholine liposomes by passive diffusion hasbeen derived by Herring et aV76, and has also been given by Nelson210. Themathematical equation describing the exponential decay of the DMA peak integralto equilibrium value is given below.- + (I - 1’)exp-{(1 .i-f)kt}[3.0]I, I° are the integrals of the DMA peak inside the liposome at time t,equilibrium, and zero time respectively. Zero time refers to time before spectralacquisition; f, is the volume ratio of the intraliposomal compartment to theextraliposomal compartment (i.e, f=V1/V0); k is the observed rate constant forDMA efflux from the liposome and t, is the efflux time in seconds.3.9.2 Facilitated diffusion.Another transport mechanism which was considered because some of theexperimental data obtained in the present study did not fit equation [3.0], is102facilitated diffusion. In facilitated diffusion, the diffusing molecule enters into someform of reversible chemical association or complexation with a carrier moleculewhich transports it across the membrane.In solution at pH 7.4, the probe molecule dimethylarsinic acid (DMA) existsas two chemical species210: the undissociated dimethylarsinic acid (DMAH), and theanionic species DMK. Both species are capable of passive diffusion in themembrane, but only DMK is likely to be transported by triorganotin cation duringfacilitated diffusion. Therefore, in this study, DMA refers to a mixture of DMAH andDMK present in solution.A proposed schematic diagram of tributyltin cation acting as a carrier is shownin Figure 3.5. In this scheme, DMA diffuses into the lipid bilayer where it associateswith the tributyltin cation to form a tributyltin-DMA complex which is mobile withinthe lipid bilayer. At the interface of the liposome and the extraliposomalcompartment, the tributyltin-DMA complex dissociates to liberate the DMK into theextraliposomal compartment.A theoretical treatment for facilitated diffusion based onHxhe adsorptionequilibria of Langmuir2 has been described by Widdas2’ to account for theplacental diffusion of glucose, and by Hall and Baker3.The theoretical treatment by Widdas2’ is further developed below fortrialkyltin cation mediated transport of DMK across a liposomal membrane.(a) DMK can associate with the tributyltin cation which is the carrier to form atributyltin-DMA complex.103Lipid bilayerIntraliposomal Extralqx)somaJcompartment compartmentDMA + (C4HSn (C4HSn + DMA—K1— kT(C4H,)SnDMA (C4HSnDMA (C4HSnDMA (C4HSnDMAk1 = k2 is the formation rate constant for (C4H9)3SnDMA.k1 = k2 is the dissociation rate constant for (C4H9)3SnDMA.= k/k is the equilibrium dissociation constant for (C4H9)3SnDMA.B = k1 1k.. is the equilibrium formation constant for (C4H9)3SnDMA.B/ is the ratio of equilibrium formation constant to equilibrium dissociationconstant of the carrier-permeant complex.K- is the partition coefficient of the carrier-permeant complex in the interfacebetween the intraliposomal compartment and the bilayer.K0 is the partition coefficient of the carrier-permeant complex in theinterface between the extraliposomal compartment and the bilayer.kT and kT are the transfer rate constants of (C4H9)3SnDMA to theextral iposomal and intral iposomal compartments respectively.Figure 3.5 Proposed mechanism of tributyltm mediated efflux of dimethylarsinate(DMA) from a liposome and the equilibria of the carrier-permeantinteractions.104(b) The tributyltin-DMA complex travels through the membrane and uponreaching the other surface dissociates and liberates the DMA into theadjacent medium. The free organotin cation returns to its original positionmay be, with a different anion in solution to start a new permeation cycle.(c) The carriers are in equilibrium with the substrate at the interfaces.(d) The carriers pass backwards and forwards across the liposomal membrane.(e) The rate of transfer of the carrier-substrate complex is much smaller than therate of formation and dissociation of the complex, and is the rate limiting step.(f) The net rate of transfer is proportional to the difference in the fraction ofsaturated carriers on both liposomal membrane interfaces.According to Langmuir211,the adsorption equilibrium at any interface can beexpressed as:13-c_____— 4) [3.1]13C÷4)-c+i4)where e is the fraction of tributyltin saturated with DMK. C, is the concentrationof DMK in solution at the interface, is the dissociation constant of the carrierpermeant complex, and B is the formation constant of the carrier-permeant complex.According to Widdas212, equation [3.11 assumes the form of the expression ofconcentration of enzyme combined with substrate in Michaelis-Menten’s equation214for enzyme kinetics (Appendix C). Therefore, Widdas212 considered , analogous tothe Michaelis-Menten’s constant. Also, Hall and Baker2’3 considered the dissociation105constant of the carrier-permeant complex as a Michaelis-Menton’s constant.The rate of disappearance of the tributyltin-DMA complex at the interfacebetween the lipid bilayer and the intraliposomal compartment (mi), is given by theexpression: -an—-+ kO [3.2]where n, refers to the number of moles of DMA in the interface between thebilayer and the intraliposomal compartment. erni, and e refer to fraction of carrierssaturated with DMK at the inside and outside interfaces of the liposomerespectively, and kT and k..T are the transfer rate constants of the tributyltin-DMAcomplex towards the outside and inside interfaces of the liposome respectively.Substituting equation [3.1] into equation [3.2], the rate of disappearance ofthe carrier-permeant complex at the inner interface (mi) becomes:j3-c j-c__________[331Tp+1prniIf the concentrations Cmi and C are small such that (i3I)C <<1, equation [3.3]becomes :-an.-+ k.TC) [3.4]13/ is the ratio of formation constant to dissociation constant for the tributyltin106DMA complex.Re-writing equation [3.41 in terms of moles of carrier-permeant complex:.S. {kT(.?L) + kT(!L.)} [5]where Vmj and V refer to the volumes of the inner and outer interfaces of theliposomes respectively, n and n refer to the moles of carrier-permeant complexin the inner and outer interfaces respectively.C. nV-fl:Vnn [3.6]Where K. is the partition coefficient of the carrier-permeant complex in the insideinterface between the liposome and the intraliposomal compartment.Therefore,Icnvm,‘mi- V [3.71Also,- -- moout oIt moTherefore,[3.8]KV— I,out107Substituting equation [3.7] and [3.8] into equation [3.5], and simplifying theresulting equation, the rate of disappearance of the carrier-permeant complex at theinner interface (rate of decrease of the DMK NMR signal inside the liposome)becomes:an V nI n a- “ Ji_J K —a + k K —at içv p T in -T [3.91If k = k.T = k, K1 = and Vmj c V, equation [3.9] becomes:an. nm. n- V kZJ- +l 1’ v0 [3.10]N, the total number of moles of DMA in both the intraliposomal and extraliposomalvolumes, and the outer and inner interfaces is given by the equation:N=nin+no,#+nmj+n,,But, n and n are very small when compared to nir. and n0 because of the verysmall volumes of the inner and outer liposomal interfaces. Therefore,n01-N-n[3.11]Substituting equation [3.111 into equation [3.10] and simplifying,108— V. k1--+ (N[3.12]ãt Lfl4,lv VUI outSimplifying equation [3.12],-_Vk13f 1 i Ni [3.13]&( + +çLet,1’ V [3.14]Substituting equation [3.14] into equation [3.13],aflflVk / N— nV+—at-vj [3.15]Re-arranging equation [3.15] and integrating,_____________— V. k--f& [3.16]UIviout,--- In In V’- —-I] - V k-1t [3.17]‘i’n V a 4)OUt109N- exp_ {v’v. k--tn4fVI- 1+f [3.20]Where f is the volume ratio of the intraliposomal to the extraliposomal volumes(f=VN0)Substituting equation [3.20] into equation [3.19] and re-arranging the resultingequation,(1+f) — N ( o 11 + f N “ 1 +f vv j7 - 112jI J - v) {(i) p f____—Iexpin outinRe-arranging equation [3.18],— —V’V-.kt[3.18][3.19]Let V’ also be:At equilibrium,r eqt -, , fin— in[3.211[3.22](n( 1+!) NVout1+f ) ckt} -, 0Therefore the expression for N in equation [3.21] becomes,110—+ (n — n1)e _{( 1 + f )kt} [3.25]The peak area of the proton resonance in ‘H NMR is directly related to the numberof particles, therefore equation [3.25]can be re-written in terms of the integral of themethyl resonance of the DMK -When there is no facilitated diffusion, there is no B or therefore, equation[3.26] reduces to equation [3.0] for passive diffusion previously derived by Herringet al’76. Equation [3.26] predicts the exponential decay of the DMK resonanceinside the liposome with time, and shows that facilitated diffusion is controlled by theratio of the formation constant to the dissociation constant B/ for the carrier-[3.23]N- eq___V Iin )Substituting equation [3.23] into equation [3.21] and simplifying,- - (n - n)exp— {( 1 + f ) - kt} [3.24]Re-arranging equation [3.24],permeant complex.1113.10 EXPERIMENTAL3.10.1 Instrumentation3.10.1.1 Nuclear magnetic resonance spectromeiry (NMR).A Bruker AM400 NMR spectrometer was used to obtain all NMR spectra.NMR facilities were provided by Professor F.G. Herring. The spectrometer wasoperated in the water suppression mode. The 5mm NMR tubes used for all theexperiments were obtained from Norell Inc., Landisville, New Jersey, U.S.A.. Theoperating parameters that were used for spectral acquisition and water suppressionare given in Appendix B.3.10.1.2 Lipid extruder and membrane filters.The lipid extruder used to produce unilamellar vesicles (liposomes) wasprovided by Professor F.G. Herring, and was purchased from Lipex BiomembranesInc., Vancouver, Canada. The 200 nm pore-sized polycarbonate filters used with theextruder were purchased from Costar Corporation, Cambridge, Massachusetts,U.S.A..3.10.1.3 UV-Visible spectrophotometry.A Shimadzu 600 spectrometer was used for all phosphorus assays. Allmeasurements were taken at 815 nm.1123.10.2 Chemicals and reagents.Tributyltin chloride was purchased from Ventron (Alfa Inorganics) Beverly,Massachusetts, U.S.A.. Monobutyltin trichioride and deuteriated 3-(trimethylsilyl)propionic acid sodium salt 2,2,3,3-d4 (TSP) were procured from Aldrich ChemicalCompany, Milwaukee, U.S.A.. Dimethylarsinic acid (DMA) was obtained fromFisher Scientific Company, Fairlawn, New Jersey, U.S.A..Tris(hydroxymethyl)aminomethane hydrochloride (tris buffer) and c-D(+)-glucosewere purchased from Sigma Chemical Company, U.S.A.. Egg phosphatidylcholine(EPC) was procured from Avanti Polar Lipids, Birmingham, Alabama, U.S.A..Solutions of the tris buffer were prepared by dissolving appropriate amounts in deionized water and adjusting the pH to 7.4 with sodium hydroxide. All solvents usedfor lipid extraction were Spectrograde. A stock solution of DMA (25 mg/mL) wasprepared in tris buffer (300 mM), and its pH was adjusted to 7.4 with sodiumhydroxide solution. The organotin compounds were freshly dissolved in tris buffer (40mM), and their pH was adjusted to 7.4 with sodium hydroxide solution if necessary.3.10.3 Preparation of large unilamellar vesicles (LUVs) from eggphosphatidyicholine (EPC) and the encapsulation of dimethylarsmic acid.A stock solution of EPC was prepared by dissolving EPC (1 g) in chloroform(10 mL). This stock solution was stored in the freezer until needed. The stocksolution (2 mL) was pipetted into a test-tube, and the solvent was evaporated offby using a gentle flow of nitrogen gas, and then the resulting paste was dried for 3113hours on a vacuum line. Multilamellar vesicles (MLVs) were then prepared byadding 300 mM tris buffer (1 mL) containing dimethylarsinic acid (25 mg/mL) ata pH of 7.4. The suspension was vortex mixed for 5 minutes, and then subjected tofive freeze-thaw cycles, according to the method of Meyer et al . The sample wasdipped in liquid nitrogen for about 2 minutes and thawed in a water bath (30 °C).The freeze-thawed vesicles were then forced by using pressure from a nitrogen tank(200-500 psi), to pass through two stacked 200 nm pore sized polycarbonate filtersin an extruder, to afford large unilamellar vesicles (LUVs). The LUVs were dividedinto two portions of about 0.4 mL, to allow duplication of each DMA effluxexperiment. For the DMA efflux experiments, the LUVs (0.4 mL) were applied ontoa Sephadex G-50 gel permeation column (1.5 cm i.d x 4 cm), pre-equilibrated in trisbuffer (40 mM, pH 7.4). Elution was achieved by the use of further 40 mM trisbuffer (pH =7.4). Upon the application of the LUVs onto the gel permeation column,timing was initiated. Only about the first 1 mL of the eluted LUVs were collected.An aliquot of the eluted LUVs (400 L) was pipetted into the NMR tube whichalready contained the following: of-D(+)-glucose (28 mg), manganese sulfate (40 jLof 30 mM), TSP (25 L of 40 mM), and tris buffer (135 L 40 mM, pH 7.4). Theuse of glucose was to control the osmotic pressure on the liposomes. The amount ofglucose added was calculated to approximately balance the osmotic pressure actingon the liposomes.The time course for the efflux of DMA from the EPC LUVs was followed byacquiring NMR spectra at appropriate time intervals, until equilibrium was reached.114The operating parameters that were used for spectral acquisition and watersuppression are given in Appendix B.The following experiments were performed on the liposomes in which DMAhad been encapsulated:(i) Efflux of encapsulated DMA from the liposome in the absence of anyorganotin compound.(ii) Efflux of encapsulated DMA from the liposome, with organotin compound(trimethyltin hydroxide or tributyltin chloride or monobutyltin trichioride)added in the extraliposomal compartment.3.10.4 Preparation of butyltin-EPC LUVs and the encapsulation of DMA.The stock EPC solution (2 mL) in chloroform was pipetted into a test tube.Also, aliquots of tributyltin chloride or monobutyltin trichioride (0.5, 1.5 or 5gImL) in chloroform (1 mL) were pipetted into the same test-tube, and vortexmixed. The chloroform was evaporated off, and the butyltin-EPC mixture dried ona vacuum line for 3 hours. DMA (1 mL of 25 mglmL solution) in tris buffer (300mM, pH 7.4) was added to the dried butyltin-EPC mixture and vortex-mixed forabout 5 minutes, to achieve the encapsulation of DMA in the butyltin-EPCmultilamellar vesicles (MLVs) that were formed. The butyltin-EPC MLVs wereforced to pass through two stacked 200 nm pore sized polycarbonate filters in theextruder, as described in Section 3.10.3 to produce large unilamellar vesicles(LUVs). The butyltin compounds possess highly hydrophobic butyl groups which115favour their incorporation into the lipid bilayer. However there will be some residualorganotin compounds in the intraliposomal compartment. Butyltin-EPC liposomes ofthe following composition were prepared:(a) 0.5jg tributyltin chloride : 0.2g EPC (TBT-EPC A)(b) 1.5ig tributyltin chloride : 0.2g EPC (TBT-EPC B)(c) 5.0ig tributyltin chloride : 0.2g EPC (TBT-EPC C)(d) 0.5/Lg monobutyltin trichioride : 0.2g EPC (MBT-EPC A)(e) 1.5.ig monobutyltin trichioride : 0.2g EPC (MBT-EPC B)The butyltin-EPC LUVs were divided into two portions of about 0.4 mL each, topermit the duplication of each DMA efflux experiment. A portion of the butyltinEPC liposomes (0.4 mL) was added onto a Sephadex G-50 gel permeation column(1.5 cm i.d x 3.0 cm) and eluted as described in Section 3.10.3. The first fraction(about 0.7 mL) of the eluted butyltin-EPC was collected. An aliquot of the elutedbutyltin-EPC liposomes (400 .iL) was quickly pipetted into the NMR tube whichalready contained glucose (28 mg), aqueous manganese sulfate solution (40 L of 30mM solution), TSP (25 L of 40 mM solution) and iris buffer(135 .iL of 40 mMsolution, pH 7.4). When the presence of butyltin chloride was desired in theextraliposomal compartment, 135 L of 16.7 J.LM solution in iris buffer of the samebutyltin chloride used to form the liposome was added. The DMA efflux experimentmonitored by using the NMR spectrometer was conducted, as described in section3.10.3 above.The following experiments were performed on the butyltin-EPC liposomes:116(i) Efflux of encapsulated DMA from the butyltin-EPC liposomes with nobutyltin chloride added into the extraliposomal compartment.(ii) Efflux of DMA from the butyltin-EPC liposomes with tributyltin chloride ormonobutyltin trichloride added into the extraliposomal compartment.3.10.5 The NMR water suppression and spectral acquisition conditions for DMAefflux from EPC and butyltin-EPC liposomes.The Bruker AM400 NMR spectrometer was operated in the water suppressionmode, which made it possible to obtain the NMR spectra of samples prepared inaqueous buffers. The water signal suppression was achieved by applying a narrowpresaturation pulse at the frequency of the water signal, followed by a broadbandexcitation pulse which was applied at the frequency of the DMA resonance, while thewater signal was still saturated.The time course for the efflux of DMA from the LUVs was followed byspectral acquisition at various time intervals. At each time interval, 48 scans werecollected and averaged. The free induction decays were acquired by using a pulsewidth of 6 milliseconds, and Fourier transformed with a line broadening of 10 Hz.The spectrometer has variable temperature capability, which was used in someexperiments.The micro-program used to operate the NMR spectrometer in the watersuppression mode with automated spectral acquisition is given in Appendix B.1173.10.6 Determination of phospholipid concentrations by phosphorus assay.3.10.6.1 Extraction of phospholipid from liposomes prior to phosphorusdetermination.Prior to determining the phosphorus concentration of the liposomes, thephospholipid (EPC) was separated from DMA by extraction into chloroform becausearsenic interferes with the subsequent phosphorus determination.The extraction was carried out according to the following procedure. TheLUVs (0.5 mL) were diluted to 1 mL with deionized water. Methanol (2.2 mL) andchloroform (1 mL) were added to the vesicles, and the mixture was vortex-mixed.Deionized water (1 mL) and chloroform (1 mL) were further added into the mixture,causing it to separate into two phases. The top phase contained methanol, water, andDMA. The bottom phase contained chloroform and the lipid.3.10.6.2 Lipid concentration determination.Lipid concentrations of the vesicles were then determined by analyzing theirphosphorus content as previously described by Fiske and Subbarow215,and Bottcheret a1216. Lipid phosphorus was converted to phosphomolybdic acid which wassubsequently reduced by the Fiske-Subbarow reagent to a blue compound which canbe measured colorimetrically. The procedure is as follows. Aliquots (25 L) of thechloroform extract of the vesicles were dispensed into test tubes and the chloroformwas gently evaporated off with a stream of nitrogen gas. Perchioric acid (7.25 mL)118was placed into each of the test tubes which was then covered with a marble ball,and placed in a metal test tube-holding block which was heated (180°C - 200°C, 1.5hours). The test tubes were cooled, and 7.0 mL ammonium molybdate reagent (0.22% wlv ammonium molybdate in 2% H2S04 wlv) and 0.6 mL Fiske-Subbarrowreagent (30g NaHSO3, ig Na2SO3 and 0.5g bis 1-amino-2-napthol-4-sulphonic acidin 200 mL water) were added into each of the test tubes. The contents of each testtube were vortex-mixed, heated for 15 minutes in a boiling water bath, cooled, andtheir absorbances measured at 815 nm. The samples were standardized againstknown concentrations of NaH2PO4 which had undergone similar chemical treatmentas the samples. Each assay was carried out in duplicate, and the average phophorusconcentration determined. The phospholipid concentration was then calculated fromthe relationship that 1 mole of EPC contains 1 mole of phophorus.3.10.7 ProcesHxng of the NMR spectra.For each experiment involving the efflux of encapsulated DMA from theliposomes, 25-30 data points collected over 11-17 hours were processed. Each datapoint, represents an average of 48 scans. The accumulated free induction decays(FID) were Fourier transformed with a line broadening of 10 Hz to produce theNMR spectra (Figure 3.6). In Figure 3.6, peak A (sharp singlet) is assigned to theDMA inside the liposome. Peak B (broad singlet) is assigned to DMA that hasdiffused out of the liposomes: the peak has been broadened and shifted by themanganese sulfate, a spectroscopic shift/broadening reagent added to the NMR tubeChemical shift (ppm)1H NMR spectra of DMA as it diffuses out of EPC liposomes. PeaksA and B are due to DMA inside and outside the liposomesrespectively. Peak D is due to iris buffer. All peaks are referenced toTSP (peak C).1196.0 50 4:0 3.0 2:0 1.o345 seconds2145 seconds12345 seconds(I)(ii)(iii)50 4.0 3.0 2:0 óoVDCAFigure 3.6120contents at the begining of the experiment. Peak D (6=2.4-4.5) is assigned to trisbuffer. All peaks were referenced to TSP (peak C). As the experiment progresses,the peak due to the DMA inside the liposome decreases while the peak due to theDMA outside the liposome increases (Figure 3.6, i, ii, iii). Either of the peaks canbe used to monitor DMA efflux from the liposomes. However, it was convenient tomonitor the decrease of the peak due to DMA remaining in the liposome becausethe peak area was easier to obtain by integration. The peak due to tris buffer wasused as an internal standard to nullify the effects of fluctuations in the instrument’soperating parameters. The peak area ratios of DMA inside the liposome to the trisbuffer were calculated and plotted as a function of time to describe the effluxbehaviour of the DMA molecules.Each data point is a combined signal from the methyl resonance of the twospecies of DMA namely; DMAH and DMK present in solution.3.10.8 Analysis and treatment of data.3. 10.8.1 Determination of rate constants and mode of permeation.The experimental data for every efflux of DMA from EPC or butyltinchloride-EPC liposomes were analyzed for passive or facilitated diffusion by usingequations [3.0] or [3.26] respectively.—+ (I — I)exp— f(1 + f )kt}[3.0]121Figure 3.7eqin1eqout= ,.eq+ (I-1)exP_{(1 +f)J.kt} [3.26]A plot of ln( j - 1eq) versus time gives a straight line with slope -(1+0k (Figure3.7) for passive diffusion, or -(1 +O(BI)k for facilitated diffusion, where f is the ratioof the internal volume to the external volume. At equilibrium, the ratio of the peakintegrals of the DMA signal inside the liposome to the signal outside the liposomecorresponds to f. Therefore,[3.27]0—2—4—6—80 200 400 600 800 1000 1200 1400Efflux time (s)Log plot for the efflux of DMA from EPC liposomes.122where and‘out are the peak integrals (area) due to DMA inside and outsidethe liposomes at equilibrium respectively, BI is the ratio of the formation constantto the dissociation constant of the carrier-permeant complex (see Figure 3.5). Amethod for estimating the ratio 8/, and k is described in Section 3.11.1. The totalvolume of reagents in the NMR tube for each experiment was 0.6 mL.Therefore,Vi,, +V0=O.6mL [3.28]A combination of equations [3.27] and [3.28] allows V1 and V0 to be calculated.Hence f in equation [3.27] is determined, and k is calculated.From a plot of equation [3.0], the intercept ln (I°jnI’jr.) is obtained. isestimated from the peak integral of DMA inside the liposome at equilibrium. Then,J0 is calculated. Equation [3.0] was fitted onto the experimental data points by aniterative procedure until convergence was obtained. This was done by using acommercially available mathematical software Sigmaplot 5.0 (Jandel Scientific). Thecalculated values of f, and k were kept constant while Im° was permitted tovariable by about ± 0.005 units about the calculated value. A good fit of equation[3.0] onto the experimental data points indicates efflux by passive diffusion, providedthe magnitude of the efflux rate constants is in the range expected for passivediffusion. This provision is necessary because in some situations (discussed in Section3.11.1), equation [3.0] can also fit data for facilitated diffusion.If the experimental data points did not fit equation [3.0J,they were analyzedfor facilitated diffusion by using equation [3.26].123At a pH of 7.4, which was the case for all experiments described in thischapter, Herring Li176 have determined that the major species of DMAP ,0H3C—As—OH+H20- H3C——O-+HO+I ICH3 pK= 6.28 CH3DMAH DMKFigure 3.8 Chemical species of dimethylarsinic acid (DMA) present at pH 7.4(pH at which all experiments reported in this thesis were conducted).permeating by passive diffusion is the undissociated acid represented as DMAH(Figure 3.8). The passive diffusion of the anionic species DMK is very slow.As the NMR signal obtained is composed of methyl resonances from bothDMAH and DMK, the calculated values for the efflux rate constant k, and thepermeation coefficient P, were corrected for the permeating species according to theequations210 below:[3.29]k -[3.30]P- aP’where a is the fraction of DMAH in solution, and is given by the relationship176a=[H} /(Ka + [H]) for a monoprotic weak acid. For DMAH176, a is 0.0593 at pH1247.4. K is the dissociation constant. k’, and P’ are the corrected values for thepermeating species DMAH. Fraction of DMK in solution is 1-a, and has a valueof 0.9407.3.10.8.2 Determination of permeability coefficients.The permeability coefficients were calculated according to equation 3.31k x Vjjimol.lipid— A [3.31]where k (Is) is the efflux rate constant, V is the trap volume of the liposome permol of phospholipid, A is area per j.tmol phospholipid, and has been calculated21°to be 1.81 x i03 cm2/jmo1 phospholipid.The lipid concentration (I.Lmol phospholipid) was determined by phosphorusassay as described in Section 3.10.6.2.1253.11 RESULTS AND DISCUSSION.3.11.lTheuseofDMAasaprobe inpermeabiity studies ofEPCliposomes, intheabsence and presence of organotin compounds in the extraliposomalaqueous compartment.To observe the effect of organotin compounds on the permeability propertiesof the liposomes, experiments were carried out by monitoring the efflux ofencapsulated dimethylarsinic acid (DMA) from liposomes in the presence andabsence of the organotin compounds added into the extraliposomal compartment. AtpH 7.4, two species of DMA namely DMAH and DMK are present in solution.The ‘H NMR spectra obtained are due to the combined proton resonances of thetwo species, therefore the graphs presented in this chapter describe the efflux ofDMA, while the tables of data are for DMAH or DMK efflux. Under conditions ofpassive diffusion, DMAH is the major species permeating out of the liposomes. Thepermeation of DMK by passive diffusion is very slow210,and is not treated in thepresent study, except in situations where it permeates by facilitated diffusion.The time course for the efflux of DMA in the absence of any organotincompound at 24°C is shown in Figure 3.9. This efflux behaviour conforms to a firstorder passive diffusion as demonstrated by fitting a curve through the data points byusing equation [3.0]. Previously, Herring Lai’76 had reported that DMA efflux fromEPC liposomes is by passive diffusion. The fit of the data points around the curvedportion of their graph is also similar to that shown in Figure 3.9. The rate constants126aIIFigure 3.9Parameters Valuea Data from ref 213t’12(s) 62±2k’(/s) (1.1 ± 0.04) x 10-2 (0.97 ± 0.15) x 10-2P’(cm Is) (1.7 ± 0.2) x 10-8a = t’, k’ , and P’ have been corrected for the major permeating specieDMAH.0.040.030.020.01 • Experimental data—Fit using eqn (3.0]7.0 x1040.000.0 1.0 2.0 3.0 4.0 5.0 6.0EFFLLIX TIME (s)Efflux of DMA from EPC liposomesin the extraliposomal compartment).Table 3.1 Efflux data for the diffusion of DMAH from EPC liposomes in theabsence of organotin compounds(organotin compounds are absent127permeation half-life and permeability coefficient for the efflux of DMAH in theabsence of the organotin compounds are shown in Table 3.1. Also, data for DMAHefflux from EPC liposomes in HEPES buffer obtained by Nelson21° are shown inTable 3.1, and are in close agreement with those obtained in this study. Thisindicates that tris buffer used in this study, or HEPES buffer do not introduce anysignificant effect in the efflux rate constants.The time course for DMA efflux in the presence of tributyltin chloride andmonobutyltin trichioride is shown in Figures 3.10 and 3.11 respectively.The permeability data for DMAH efflux in the presence of tributyltin chloride in theextraliposomal aqueous compartment are shown in Table 3.2.A comparison of the efflux data of DMAH from EPC liposomes in theabsence of any organotin compound (Table 3.1), and its efflux data in the presenceTable 3.2 Effect of 33.2 M tributyltin chloride on the efflux of DMAIIfrom EPC liposomes.Parameter Valuet’ (if diffusion is passive) 25 ± 1k’(Is) (if diffusion is passive) (2.8 ± 0.1) x 10P’ (cm/s) (if diffusion is passive) (4.9 ± 1.0) x 1080.020128IIFigure 3.100.0150.0100.005 -0.000• Experimental data—Fit using eqn [3.0]Fit using eqn [3.26]I I I I I I0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 x 10EFFLUX TIME (s)Time course for the efflux of DMA from EPC liposomes (tributyltinchloride present in the extraliposomal compartment).—Iz0.005- -< 0.000 I I I0.0 0.5 10 1.5 xEFFLUX TIME (s)Figure 3.11 Time course for the efflux of DMA from EPC liposomes (monobutyltintrichioride present in the exiraliposomal compartment).-• Experimental dataFit using eqn 13.0]129of tributyltin chloride (Table 3.2) shows that in the presence of tributyltin chloride,the permeation half-life t’ of DMAH efflux becomes about 2.5 times smaller thanin its absence, indicating increased rate of permeation. The permeability coefficientis also about 2.9 times greater than in the absence of tributyltin chloride. The effluxrate constant k’ also increased by more than twice. This increased permeability ofDMAH observed in the presence of tributyltin chloride suggests that some propertiesof the EPC liposomes have been changed by the tributyltin chloride. According toHeywood et a!209, the tributyltin cation causes membrane disruption and rupture(lysis) of EPC liposomes. Such rupture or pore formation in the liposomal bilayerwould result in increased permeability to permeants.The increased permeability of the EPC liposomes to dimethylarsinic acid inthe presence of tributyltin chloride could also arise if the tributyltin cation acted asa carrier, and mediated the transport of DMK by facilitated diffusion. The abilityof the tributyltin cation to facilitate the diffusion of CF and OW has been reportedby Selwyn15’208 and Tosteson’6.The equations [3.0] for passive diffusion and [3.26] for facilitated diffusion,I, -1’ + (1 —1)exp -{(1+f)kt}[3.0]-+ (l — 1 )exP_{(1 + f)..P_kt} [3.26]used in this study for fitting curves onto the observed experimental data, are notcapable of distinguishing between situations where there is a 100% passive diffusion130or 100% facilitated diffusion, unless the magnitudes of the expected rate constantsare known. This is because when there is 100% passive diffusion or facilitateddiffusion, a plot of ln(It1 4el.r) versus t, gives a straight line with slope (1+0k forpassive diffusion or (1 +f)(Bhk)k for facilitated diffusion. Each of the constants BIband k for facilitated diffusion cannot be calculated separately, instead they areincorporated into each other as one parameter. The combined value of (i3/)k ofequation [3.26] is equivalent to k of equation [3.0]. Under this condition, any curvegenerated by using either equation [3.0] or [3.26] should fit the experimental data(Figure 3.10).A method for obtaining the value of the constant 13/i when efflux takes placeby a mixture of passive diffusion and facilitated diffusion is described later in thissection.The time course for the efflux of DMA in the presence of monobutyltintrichioride is shown in Figure 3.11 above. The efflux of DMAH is by passivediffusion, but the rate of efflux has become slower (Table 3.3). The values of therate constant and permeability coefficient shown in Table 3.3 are smaller than theirvalues when no organotin compound was present (Table 3.1). The permeation halflife of DMAH is about 3 times larger, indicating retarded permeation. Also, thepermeability coefficient is decreased by about a factor of 4.3.The effect of externally added monobutyltin trichioride on the EPC liposomesis to decrease their permeability. A probable mechanism for this behavior is thatmonobutyltin trichloride permeates into the lipid bilayer and causes a decrease in the131membrane fluidity. Any compound capable of decreasing membrane fluidity such ascholesterol217 and o-tocopherol218 would decrease permeability. Such decrease in themembrane fluidity caused by dibutyltin dichioride has been observed onphosphatidylinositol 4-monophosphate and phosphatidylinositol 4,5-diphosphateTable 3.3 Effect of 33.2 1iM monobutyltin irichioride on the efflux ofDMAH from EPC liposomes.Parameters Valuetlk (s) 188 ± 6k’ (Is) (3.7 ± 0.1) x i0P’ (cm/s) (4.0 ± 1.2) x i0vesicles2’920.Unfortunately, apart from the present study, there are no other reportsof the interaction of monobutyltin trichloride with liposomal membranes. However,from the data presented in Tables 3.2 and 3.3, it seems that the monobutyltinspecies, unlike the tributyltin species, neither have the capability to induce membranedisruption nor act as carrier. Hence, the retarded permeation of DMAH.Monobutyltin is a degradation product of tributyltin in the environment andthe scheme is tributyltin -. dibutyltin -‘ monobutyltin —‘ inorganic tin. These productsare progressively less toxic to life perhaps because debutylation leads to products lesscapable of causing membrane disruption.132The efflux of DMA across EPC liposomes in the presence of trimethyltinhydroxide (32.3 tiM) in the extraliposomal compartment was also studied. Analysisof this permeation behaviour by plotting ln(It1- Ifr) against t, of equation [3.0]or [3.26] gave two straight lines of different slopes (Figure 3.12, slopes A and C).This behaviour is not predicted by equation [3.0], if permeation is by passivediffusion.i— + (1 — 1)exp —((1 +f)kt}[3.0]- Jeq+ (I - 1)exp _{(1 + f )!kt} [3.26]If slope A (Figure 3.12) is considered a region of facilitated diffusion, the parameter(fi/)k of equation [3.26] is calculated. When the value of (BI)k was used inequation [3.26] to fit the experimental data, only the data points from the early andvery late parts of the efflux experiment fitted (Figure 3.13 solid curve). Dataobtained at intermediate efflux times did not fit.If slope C (Figure 3.12) is considered a region of passive diffusion, the effluxrate constant k, for DMAH is calculated. When k was used in equation [3.0] to fitthe experimental data, there was a close fit for data obtained at intermediate andlate parts of the efflux (Figure 3.13 broken line). Towards equilibrium, equations[3.0] and [3.26] gave a close fit. A possible explanation for this phenomenon is thatslope A (Figure 3.12) is a region dominated by facilitated diffusion of DMKmediated by trimethyltin cation, while slope C is a region dominated by passive133-a—5.0I.Figure 3.12ABCA= Region of facilitateddiffusion.B = Mixture of facilitated& passive diffusion.C = Region of passivediffusion.(33.2 M IrimethyltinI - I0 1000 2000 3000Efflux time (s)Log plot of DMA efflux from EPChydroxide present in extraliposomalliposomesvolume).0,030.02-- Fit usingeqn [3.0]—Fit usingeqn [3.26]— 0.01o -‘- Fit usingeqn [3.31]0.000 5000 10000 15000 20000 25000EFFLIJX TIME (s)Figure 3.13 Time course for DMA efflux from EPC liposomes (33.2 tMtrimethyltin hydroxide present in extraliposomal volume).134diffusion of DMAH and DMK. The passive diffusion of DMK is very slow210.SlopeB (Figure 3.12) is a region dominated by a mixture of facilitated and passivediffusion. Thus, facilitated diffusion sets in at the early part of the efflux, andgradually gives way to passive diffusion as the efflux progresses. Assuming that theDMA is the major species permeating by facilitated diffusion while DMAH is themajor species permeating by passive diffusion, the rate constant obtained from slopeC (Figure 3.12) should be divided by 0.9407; the fraction of DMA (1-at) present atpH 7.4 (Section 3.10.8.1), to obtain the rate constant k’ for the passive efflux ofDMA. Then, the parameter 8/ for facilitated diffusion of DMA can be calculatedfrom slope A (Figure 3.12) by using the rate constant k’ (for passive diffusion ofDMK) calculated from slope C.The calculated parameters k’ (DMK), k (DMAH), B/, and l aresubstituted into equation [3.31] which is a combination of equations [3.0] and[3.26], modified by introducing the parameters M and N. Equation 3.31 was fittedonto the experimental data by keeping every other parameter except M and Nconstant (Figure 3.13, dotted line).-M(1+(I-1)exp-((1+t)kt}) + N(1+(I1 .[3.31](M is the percentage contribution of passive diffusion to efflux, while N is thepercentage contribution of facilitated diffusion to efflux). The efflux parameters forthis experiment are shown in Table 3.4.135Similar increased flux of chloride ions across liposomal membranes, mediatedby trimethyltin cation as carrier has been reported by Selwyn15.Table 3.4 Data for the efflux of DMA from EPCpresence of 33.2 jcM trimethyltin hydroxide.liposomes in theParameter Valuek’ DMAH (Is)k’ DMK (Is)BI (experimental)aB/ (curve fit)”N (% facilitated diffusion)bM (% passive diffusion)L(2.5 ± 0.5) x(1.6 ± 0.3) x2.5 ± 0.52.2 ± 1.066 ± 2135 ± 21a= Calculated from experimental datab=obtained from the curve fitting result.3.11.2 Effect of organotin concentration on the efflux of DMAThe effect of tributyltin chloride concentration on the efflux of DMAH acrossEPC liposomes is shown in Table 3.5. As the concentration of tributyltin chloridein the extraliposomal compartment is increased from 0 to 8.3 pM, there is an initialdecrease in the efflux rate constant and the permeability coefficient, followed by an136increase as the tributyltin concentration is raised from 8.3 to 33.2 tM. The valuesfor the efflux half-life t’ also change in accordance with the changes in thepermeability coefficients and the efflux rate constants, by becoming smaller as thepermeability of the liposome increases.The increase in the permeability coefficients, efflux rate constants, and thedecrease in the efflux half-lives indicates that either the liposomal membrane hasbecome more permeable to DMAH or that the tributyltin cation is facilitating theefflux of DMK.The effect of tributyltin chloride on the permeation of DMAH isconcentration dependent. A plot of efflux rate constant versus tributyltin chlorideTable 3.5 Effect of tributyltm chloride concentration on DMAII effluxaCone. P’ (cm/s) t’ (s) k’(Is)(I.LM) x 10 x 1020.0 (1.7 ± 0.2) 62 ± 2 (1.1 ± 0.04)8.3 (1.3 ± 0.1) 85 ± 11 (0.8 ± 0.1)16.7 (1.6 ± 0.1) 45 ± 0.4 (1.5 ± 0.02)33.2 (4.9 ± 1.0) 25 ± 1 (2.8 ± 0.1)a=It could not be determined if increased efflux was by passive orfacilitated diffusion.137—5 0 5 10 15 20 25 30 35M (C4H9)3SnC1Effect of tributyltin chloride concentration on the permeability of EPCliposomes (tributyltm chloride was added into the extraliposomalcompartment).2 x 10-24 x 103 x 102 x 1021 x 1020Figure 3.141 x 1025 x io-0—5 0 5 10 15 20 25 30 35 40JLM (C4H9)SnC13Figure 3.15 Effect of monobutyltin trichloride on the permeability of EPCliposomes (monobutyltin trichioride was added in the extraliposomalcompartment).138concentration shows a linear relationship (Figure 3.14) described by the equationY=8.01 x i0’ X + 1.46 x iti, and a regression coefficient of 0.9990. The datapoint corresponding to zero concentration of tributyltin chloride does not fall on theregression line. This is probably because tributyltin chloride modified the propertiesof the liposomes. Therefore, the data obtained at zero tributyltin chlorideconcentration, and the other data points effectively belong to different types ofliposomes. Alternatively, non-linearity could also result if different modes of transportof DMA exist between the liposomes which are in contact with tributyltin chloride,and those not in contact with it.Table 3.6 Effect of monobutyltin trichioride concentration on the efflux ofDMAH.Conc P’ (cm/s) t’ (s) k’(Is)(jiM) x i0 x i00 (17.3 ± 0.2) 64 ± 0.5 (111.0 ± 0.04)16.7 (2.8 ± 0.4) 188 ± 6 (3.7 ± 0.1)33.2 (2.8 ± 0.1) 188 ± 6 (3.7 ± 0.1)The effect of monobutyltin trichloride concentration on the permeation ofDMA across EPC liposomes is shown graphically in Figure 3.15 , while thepermeability data are shown in Table 3.6. As the concentration of monobutyltin139trichioride is increased from 0 to 16.7 jtM, there is a large decrease in thepermeability coefficients and the rate constants. The permeation half-life increasedby about thirty fold, indicating a very retarded permeability of the liposomal bilayer.No further change in the permeation parameters was observed as the concentrationof monobutyltin trichioride was further increased.3.11.3 Efflux of DMA from tributyltin chloride-EPC liposomes (with tributyltinchloride absent in the extraliposomal compartment).The efflux of DMA from liposomes composed of a mixture of tributyltinchloride (TBT) and egg phosphatidylchloine (EPC) was studied to establish thepermeability properties of these model membranes. In these experiments notributyltin chloride was added to the extraliposomal compartment. The liposomeswere prepared as described in Section 3.10.4 and are designated as TBT-EPCliposomes. The composition of the different TBT-EPC liposomes that were studiedis as follows:(a) TBT-EPC A (0.5 g tributyltin chloride : 0.2 g EPC)(b) TBT-EPC B (1.5 ig tributyltin chloride : 0.2 g EPC)(c) TBT-EPC C (5.0 ig tributyltin chloride : 0.2 g EPC)The permeability properties of these liposomes were studied by measuring theefflux of encapsulated DMA from these liposomes. As an example, the efflux ofDMA from TBT-EPC C liposomes is described. Analysis of this efflux experimentby plotting lfl(It - eq r) of equations [3.0] or [3.26] as described in Section 3.11.1Figure 3.160 10000 20000 30000 40000 50000 60000EFFLUX TiME (s)• Experimentaldata--. Fit usingeqn [3.0]— Fit usingeqn [3.26]Fit usingeqn [3.31]140—5—6—7—sA= Region of facilitateddiffusion.Mixture of facilitated& passive diffusion.Region of passivediffusion.0 5000 10000 15000 20000 25000Efflux time (s)Log plot of DMA efflux from TBT-EPC C liposomes.0030.020.010.00Figure 3.17 Time course for DMA efflux from TBT-EPC C liposomes.141shows two lines with different slopes (Figure 3.16), which indicates that the efflux ofDMA from TBT-EPC C is by a mixture of passive diffusion and facilitated diffusion.Facilitated diffusion is the major mode of DMA transport at the early stages of theefflux (Figure 3.17 solid line), while passive diffusion of DMAH dominates at thelater stages (Figure 3.17 broken line). At intermediate times, a mixture of facilitateddiffusion and passive diffusion dominate the efflux. The observed efflux behavior istherefore better described by fitting equation [3.31](a combination of equations [3.0]and [3.26], Section 3.11.1) onto the experimental data (Figure 3.17, dotted line).Table 3.7 Diffusion parameters for the efflux of DMA from tributyltin-EPCliposomes by a mixture of passive and facilitated diffusiona.Liposome k’ (DMK) k’ (DMAH) 8/ %x i(i (Is) x iO (Is) facilitated passivediffusion diffusionTBT-EPC A 2.0 3.1 2.2 66 34TBT-EPC B 1.4 2.2 1.8 58 42TBT-EPC C 1.1 1.8 2.2 54 46a=Tributyltin chloride was not added into the extraliposomal compartment.Mean value BI = 2.1 ± 0.2142The efflux of DMA from TBT-EPC A and TBT-EPC B, can also be accountedfor by a mixture of facilitated diffusion and passive diffusion. The diffusion constants,and the percentage contributions of facilitated and passive diffusion to permeationare shown in Table 3.7.As the concentration of tributyltin chloride in the liposome is increased,facilitated diffusion decreases, while passive diffusion increases (Figure 3.18).70Figure 3.18 Contribution of passive and facilitated diffusion to the efflux of DMAfrom TBT-EPC liposomes of different tributyltm chloride composition., 60i50I4030— Facilitated diffusionPassive diffusion0 1 2 3 4 5 6tg tributyltm chloride : O.2g EPC143When tributyltin chloride (16.7 tiM) was added into the extraliposomalcompartment of TBT-EPC B liposomes, the plot of 1n(It- I ) versus t, for DMAefflux of either equation [3.0] or equation [3.26] gave one straight line of uniformslope (Figure 3.19).— 0- I I I1 0 500 1000 1500 2000 2500 3000— Efflux time (s)Figure 3.19 Log plot for efflux of DMA from TBT-EPC B liposomes when 16.7MMtributyltin chloride is present in the extraliposomal compartment.144Thus, it seems that when tributyltin chloride is added into the extraliposomalcompartment, dissociation of the tributyltin-DMA complex at the interface betweenthe bilayer and the extraliposomal compartment is no longer favourable. Theequilibrium for dissociation is shifted to the left thereby suppressing the release ofthe complexed DMK. If the tributyltin-DMA complex does not dissociate at theinterface of the extraliposomal compartment, there will be no more free tributyltincations to sustain the facilitated transfer of DMK, therefore the facilitated diffusionof DMK ceases. Under these conditions, the passive diffusion of DMAH dominates.__0.020I0.0150.010• Experimentaldata0.005—Fit usingeqn [3.0]0.0000 5000 10000 15000 20000 25000 30000EFFLUX TIME (s)Figure 3.20 Time course for efflux of DMA from ThT-EPC B liposomes when 16.7M tributyltin chloride is present in the extraliposomal compartment.145The time course for the efflux of DMA from TBT-EPC B liposome in thepresence of tributyltin chloride (16.7 JLM) added into the extraliposomalcompartment is shown in Figure 3.20. The efflux behavior is described by equation[3.0] for passive diffusion.The parameters for this efflux are shown in Table 3.8. The rate constant forthe efflux of DMAH from TBT-EPC B in the presence of externally added tributyltinchloride is 1.8 x 10-2 /s (Table 3.8), while in the absence of externally addedtributyltin chloride, it is 2.2 x i0 Is (Table 3.7). The observed difference in thetwo situations is attributed to the unequal concentrations of tributyltin chlorideinvolved in the experiments.Table 3.8 Parameters for the efflux of DMAH from TBT-EPC B liposomes in thepresence of externally added tributyltin chloride (16 SM).Parameter Valuek’ (1.8 ± 0.1) x 10-2 (Is)p’ (2.8 ± 0.5) x 10-8 (cm/s)3.11.4 Efflux of DMA from monobutyltin trichloride-EPC liposomes.Monobutyltin trichloride-EPC liposomes designated MBT-EPC liposomesprepared as described in Section 3.10.4 were used to study the permeation of DMA.146MBT-EPC liposomes having the following compositions were studied:(a) MBT-EPC A (0.5 j.g monobutyltin trichioride : 0.2 g EPC)(b) MBT-EPC B (1.5 g monobutyltin trichioride : 0.2 g EPC)The efflux of DMA from MBT-EPC B with no monobutyltin trichioride in theextraliposomal aqueous compartment was studied. The time course for this efflux isshown in Figure 3.22. Analysis of the experimental data points by plotting 1n(It -) versus t, of either equation [3.0] or equation [3.26] gave only a straight lineof uniform slope (Figure 3.21). This indicates that only one mode of diffusion isinvolved. The efflux parameters for DMAH, assuming passive diffusion is shown inTable 3.9.The efflux of DMA from MBT-EPC B in the presence of externally addedmonobutyltin trichioride (16.7 .iM) was studied. Also, only one mode of DMA effluxwas found. The efflux parameters for DMAH , assuming passive diffusion is shownin Table 3.10.It seemed reasonable to consider the efflux of DMAH from MBT-EPCliposomes to be by passive diffusion either in the presence or absence ofmonobutyltin trichioride in the extraliposomal compartment, because a comparisonof the efflux parameters in Tables 3.9 and 3.10, with the efflux parameters for an“EPC only” liposome (Table 3.1), shows that the permeability coefficient for theMBT-EPC liposomes either in the presence or absence of externally addedmonobutyltin trichioride is the same as the permeability coefficient for the efflux ofI147-4.0I-4.2—4.41 —4.6I—4.81 -5.0Figure 3.210 500 1000 1500Efflux time (s)Log plot of DMA efflux from MBT-EPC B liposomes.• Experimentaldata—Fit usingeqn [3.0]0.020.010.000 10000 20000 30000EFFLUX TIME (s)40000Figure 3.22 Time course for DMA efflux from MBT-EPC B liposomes.148Table 3.9 Permeability data for efflux of DMAII from MBT-EPC B (1.5 gbutyltin trichloride: 2 g EPC) liposomes (with monobutyltintricbloride absent in the extraliposomal compartment).Parameter Valuek’(Is) (8.3 ± 0.4) x i0t’(s) 84±4P’(cm/s) (1.7 ± 0.1) x 10-8Table 3.10 Permeability data for efflux of DMAH from MBT-EPC B (l.5jcgmonobutyltin trichioride: 0.2g EPC) liposomes with monobutyltinIrichloride present in the extraliposomal compartment.Parameter Valuek’(/s) (9.1 ± 0.5) x i0t’(s) 76±4P’(cm/s) (1.7 ± 0.1) x 10.8DMAH from “EPC only” liposomes. The efflux rate constants in Tables 3.1, 3.9 and3.10, are close. Therefore, monobutyltin trichioride does not have the ability to actas carrier for DMK.149It seems that the incorporation of monobutyltin trichioride into the lipidbilayer of EPC liposomes has no effect on its membrane permeability, but themembrane permeability is greatly retarded if the monobutyltin trichioride is addedexternally into the extraliposomal compartment (Section 3.11. 1,Table 3.3).3.11.5 Effect of the butyltin chloride concentrations of the liposome on permeabilityproperties of TBT-EPC and MBT-EPC liposomes.The experiments reported in this section were conducted with tributyltinchloride (16.7 jIM) added into the extraliposomal compartment of TBT-EPCliposomes because under these conditions, passive diffusion of DMA is induced.Monobutyltin trichioride (16.7 M) was also spiked into the extraliposomalcompartment of MBT-EPC liposomes to maintain similar experimental conditionswith the MBT-EPC liposomes.The efflux of encapsulated DMA from butyltin-EPC liposomes was monitoredat 24 °C. Studies were conducted by using TBT-EPC A, TBT-EPC B, MBT-EPC A,and MBT-EPC B liposomes.The permeability data for the efflux of DMAH from TBT-EPC liposomes areshown in Table 3.11. As the tributyltin chloride concentration of the liposome isincreased on going from TBT-EPC A to TBT-EPC B, the permeability of theliposomes to DMAH also increases by about a factor of 1.6 (Table 3.11).The permeability data for MBT-EPC liposomes (with monobutyltin chloridepresent in the extraliposomal volume) are shown in Table 3.12. The permeability150coefficients, permeation half-lives, and rate constants show very little variation withincrease in the monobutyltin trichioride composition of the liposome.Table 3.11 Effect of tributyltin chloride concentration of TBT-EPC liposomes onpermeability (tributykin chloride solution was also added to theextraliposomal volume.Liposome P’ (cmls) t’js) k’(Is)x 10-8 x 10-2TBT-EPC A (1.7 ± 0.2) 57 ± 4 (1.1 ± 0.1)TBT-EPC B (2.8 ± 0.5) 39 ± 1 (1.8 ± 0.1)Table 3.12 Effect of monobutyltin trichioride concentration of MBT-EPCliposomes on permeability (monobutyltin trichioride was alsoadded to the extraliposomal volume).Liposome P’ (cm/s) t’%(s) k’(Is)x iO x i0MBT-EPC A (4.9 ± 0.8) 195 ± 7 (3.6 ± 0.1)MBT-EPC B (4.4 ± 0.3) 218 ± 19 (3.2 ± 0.2)1513.11.6 Effect of temperature on the permeability of organotin-EPC liposomes.The effect of temperature on the permeability of TBT-EPC A and TBT-EPCB liposomes is shown in Tables 3.13 and 3.14 respectively, while the effect ofTable 3.13 Effect of temperature on the permeability properties of TBT-EPC Aliposomes.Temperature °C P’ (cmls) k’(Is)x 10-8 x 10.224 (1.7 ± 0.2) (1.2 ± 0.1)28 (3.1 ± 0.2) (2.3 ± 0.1)32 (3.4 ± 1.0) (4.7 ± 0.1)Table 3.14 Effect of temperature on the permeability properties of TBT-EPC B(1.5 jg iributyltin chloride O.2g EPC) liposomes.• Temperature °C P’ (cm/s) k’ (Is)x 10.8 x 10-224 (2.8 ± 0.5) (1.8 ± 0.1)28 (4.0 ± 0.03) (3.5 ± 0.06)32 (4.9 ± 0.5) (5.4 ± 1.3)152Table 3.15 Effect of temperature on the permeability properties of MBTEPC A (0.5 jig butyltin Irichionde 0.2 g EPC) liposomes.Temperature P’ (cmls) k’(/s)(°C) x i0 x i024 (4.9± 0.8) (3.6±0.1)28 (11.9 ± 1.0) (7.0 ± 0.1)32 (15.1 ± 1.2) (14.9 ± 0.6)Table 3.16 Effect of temperature on the permeability properties of MBT-EPC B(1.5 jig monobutyltin trichioride : 0.2g EPC) liposomes.Temperature P’(cm/s) k’ (Is)(°C) x i0 x 10-224 (4.4 ± 0.4) (5.4 ± 1.3)28 (8.7 ± 0.1) (7.6 ± 1.1)32 (15.8 ± 1.0) (1.1 ± 0.1)temperature on MBT-EPC A and MBT-EPC B liposomes is shown in Figures 3.15and 3.16 respectively.As the temperature is increased, the permeability of both TBT-EPC and153MBT-EPC liposomes to DMAH also increases. Generally, permeation rates increasewith increase in temperature. This is due to either increased partitioning of thepermeant into the lipophilic bilayer or the increased ease of diffusion through theliposomal bilayer as temperature increases.3.11.7 Activation energies for the permeation of butyltin chloride-EPCliposomes.The activation energies for the efflux of DMAH from liposomes of variousbutyltin chloride/EPC compositions were determined by using the Arrheniusequation: -P — AemTorInP----÷1nARTA plot of In P against lIT gives a straight line from which the activation energy canbe calculated (Figures 3.23 and 3.24 for TBT-EPC and MBT-EPC liposomesrespectively). The activation energies are shown in Table 3.17 for tributyltinchloride-EPC liposomes and Table 3.18 for monobutyltin trichloride-EPC liposomes.For the TBT-EPC liposomes, the activation energy for the permeation ofDMAFI decreases with increasing tributyltin chloride concentration in the liposome.When compared to the activation energy for the efflux of DMAH from an “EPConly” liposome (86 ± 20 kJ/mol)210, tributyltin chloride reduced the activationenergy required for DMAH efflux.154-16.0-16.5-17.0-17.5-18.0(l/T x3.40Figure 3.23 Arrhenius plot for DMAH efflux from TBT-EPC B liposomes.-16.0-17.0-18.0-19.0-20.0(l/T x 10) °K3.25 3.30 3.353.28 3.32 3.36Figure 3.24 Arrhenius plot for DMAH efflux from MBT-EPC B liposomes.155Table 3.17 Effect of tributyltin chloride content of liposome on the activationenergy for efflux of DMAI{ from TBT-EPC liposomes.Liposome Activation energy (kJ/mol)TBT-EPC A 64.4TBT-EPC B 52.3Table 3.18 Effect of monobutyltin trichloride content of liposome on the activationenergy for efflux of DMAI{ from MBT-EPC liposomes.Liposome Activation energy (kJ/mol)MBT-EPC A 106.8MBT-EPC B 121.5Cohen221, has described the activation energy for a permeant to comprise thefollowing: -(i) adsorption of the solute at the lipid membrane/water interphase(ii) dehydration of the solute(iii) diffusion through the hydrocarbon chain (lipid bilayer)156Thus the activation energy for permeation of a solute should increase as itsability to form hydrogen bonds increases. This is simply related to the number ofhydrogen bonds the permeant has to break before it diffuses across the hydrophobichydrocarbon chains of the lipid bilayer. If the permeating probe moleculedimethylarsinic acid (DMAH) is kept constant, the contribution of the dehydrationstep to the activation energy should be constant for both TBT-EPC and MBT-EPCliposomes. Therefore, the observed difference in activation energies for the two typesof liposomes must be due to either the effect of the organotin compounds on theadsorption of DMAH at the lipid/water interphase, or the effect on diffusion throughthe liposomal lipid bilayer. The low activation energy observed for the permeationof DMAH across tributyltin chloride-EPC liposomes (Table 3.17) supports theargument that tributyltin chloride modified the liposomal membrane.For the MBT-EPC liposomes, the activation energy for the permeation ofDMAH increases as the concentration of monobutyltin trichloride in the liposomesincreases (Table 3.18). This observation indicates that it became more difficult forthe DMAH molecules to diffuse across the lipid bilayer and is further evidence thatthe monobutyltin cation is neither capable of inducing pore formation on theliposomes nor able to act as a carrier for DMA.The pre-exponential factors of the Arrhneius equation are shown in Tables3.19 and 3.20 for TBT-EPC and MBT-EPC liposomes respectively. According toCohen221, and De Gier et a1202, the pre-exponential factor is related to the molar157Table 3.19 Arrhenius pre-exponential factor for TBT-EPC liposomesLiposome Pre-exponential factor (Is)x 10-8TBT-EPC A (2.8 ± 0.5)TBT-EPC B (4.0 ± 0.4)Table 3.20 Arrhenius pre-exponential factor for monobutyltin MBT-EPC.Liposome Pre-exponential factor (Is)x 10-8MBT-EPC A (1.1 ± 0.3)MBT-EPC B (1.0 ± 0.4)entropy change of the permeation process by the equation:ASIn A - constant + —Rwhere A is the pre-exponential function, AS is the molar entropy change and R is themolar gas constant.The values of the pre-exponential factors obtained for the TBT-EPC andMBT-EPC liposomes are different from each other, but are fairly constant for each158type of liposome. Therefore it seems that entropy for the permeation process isdifferent for both the TBT-EPC and MBT-EPC liposomes, but remains constant foreach type of liposome irrespective of the butyltin chloride concentration of theliposome. The slight variation in the values of the pre-exponential factors shown inTable 3.19 for the TBT-EPC liposomes may be attributed to experimental errors.The observed differences in the values of activation energies and preexponential factors for the TBT-EPC and MBT-EPC liposomes may indicate thatthese butyltin chlorides act on the model membrane by different mechanisms.Cohen221 has shown that the magnitude of the activation energy is related tothe physical state of the hydrocarbon chains in the lipid bilayer. Thus, as the amountof cholesterol content of the vesicle is increased, the activation energy for itspermeation also increases221.The results of the present study and those of Cohen221 demonstrate that thecomposition of the liposomes contributes very significantly to the magnitude ofactivation energy. This is contrary to the report by De Gier Li202 that activationenergy is solely determined by the capability of the permeating molecules to beinvolved in hydrogen bonding.3.11.8 Relevance of this NMR study to the environmental toxicity of butyltincompounds.A number of chemical reactions of the trialkyltin compounds with otherorganic molecules of biological relevance have been reported219’22. Trialkyltin159compounds derange mitochondrial function by discharging a hydroxyl-chioridegradient across the membranes, and by inhibiting ATP synthesis18.They also causeswelling and disruption of the mitochondrial membranes18,and the rupture of humanred blood cells223.Dialkyltin compounds react with enzymes possessing thiol groups18. Thebiochemical effect of this is an interference with a-keto acid oxidation18,while themono-organotin compounds do not show any significant toxicity.The effect of butyltin compounds on membranes has not been extensivelystudied. Early studies by Selwyn et al’5’208,Tosteson and Weith16,Motais ailV7 showthat tributyltin chloride and trimethyltin chloride can mediate chloride-hydroxideexchange across mitochondrial membrane and model cell membranes, whiletripropyltin chloride mediates chloride-chloride exchange across mitochondrialmembranes. The present study clearly shows that tributyltin chloride andmonobutyltin trichioride exert different and opposite effects on the model cellmembranes. Tributyltin chloride makes the model membranes more permeable whilemonobutyltin trichioride makes them less leaky. Since monobutyltin trichioride is byfar less toxic than tributyltin chloride, the observed decrease in membranepermeability is likely a phenomenon that leads to reduced toxicity.The present study concludes that tributyltin and trimethyltin cations are ableto function as mobile carriers for dimethylarsinate while monobutyltin cation lacksthis ability.160CHAPTER 4HYDRIDE GENERATION METHODS OF ATOMIC ABSORPTIONSPECTROPHOTOMERY FOR TOTAL TIN DETERMINATION.4.1 INTRODUCTION.With the advent of organotin pollution in the marine environment, manyworkers in the field of environmental analysis have devoted their energies to thedetection and quantitation of the more toxic organotin species. The determinationof the total tin content in marine samples has been largely neglected. Consequentlyanother objective of the present study was to provide information on the total tincontent of some marine animals in British Columbia, Canada.Determination of total tin content in environmental samples is usuallyaccomplished by the use of atomic absorption spectrophotometry (AAS). Methodsof sample preparation, prior to total tin determination, usually involve the extractionof the tin compounds into organic solvents by the use of complexing agents, or thedigestion of the samples with mineral acids, to convert the various forms of tin toinorganic tin. The total tin content of the digested sample or the organic extract canthen be determined either directly by the use of conventional flame AAS224’56and graphite furnace atomic absorption spectrophotometry (GFAAS), or byconversion to volatile derivatives such as inorganic tin hydride which can be analyzedby hydride generation-atomic absorption spectrophotometry (FIG-AAS)227’8 orhydride generation-graphite furnace atomic absorption spectrophotometry (HG161GFAAS)229. The HG-GFAAS method has also been used in the analysis of thefollowing elements; bismuth°, antimony1 and selenium2.Conversion of tin compounds in environmental samples to inorganic tinhydride is usually preferred over direct determination, because the analyte isremoved from the matrix of the digested sample, thereby minimizing matrixinterferences during the HG-AAS or HG-GFAAS analysis.In this study, two methods of hydride generation atomic absorptionspectrophotometry were optimized and used for total tin determination in marineanimals: one based on continuous hydride generation atomic absorptionspectrophotometry (HG-AAS) and the other on batch hydride generation-graphitefurnace atomic absorption spectrophotometry (HG-GFAAS).The continuous hydride generation method (HG-AAS) utilizes the hydridegenerator previously reported by Cullen and Dodd233 (Fig 4.1), for use in thedetermination of arsenic. Atomization of tin compounds was achieved inside a quartzcell, which was heated by the air-acetylene flame of the atomic absorptionspectrophotometer.The batch hydride generation method (HG-GFAAS) involved the in situgeneration of tin hydride which was then trapped, or adsorbed onto a graphitefurnace tube according to the method of Sturgeon eta1229.The graphite furnace tubeserved as a preconcentration device and also enabled high atomization temperaturesto be reached.The overall reaction for the production of tin(IV) hydride (stannane) is given162below;Sn4+4NaBH4HCl+l62O-. SnH4+4NaCZ+4H3B012H+4304.2 EXPERIMENTAL.4.2.1 Instrumentation.4.2.1.1 Continuous hydride generation atomic absorption spectrophotometry(HG-AAS).The continuous hydride generator employed in this study was a home builtglass apparatus described previously by Cullen and Dodd233 (Fig 4.1) for arsenicdetermination. The operation of this hydride generator is similar to the type reportedby Vijan and Chan22T and Subramanian228 for total tin determination. The hydridegenerator consisted of a 20 turn reaction glass coil (A, in Figure 4.1) connected toa gas-liquid separator (B, in Figure 4.1) by Teflon® tubing. Reagents were pumpedinto the glass reaction coil by means of a peristaltic pump (Gilson, MiddletonWisconsin, U.S.A.). The generated tin hydride was carried by a flow of nitrogen viaa Teflon® tubing, into an open-ended T-shaped quartz cell which was heated by theair-acetylene flame of the atomic absorption spectrophotometer. The light from thetin hollow cathode lamp, and the deuterium background corrector were aligned topass through the T-shaped quartz cell positioned in the optical path of the atomicabsorption spectrophotometer. The atomic absorption spectrophotometer is aVarian 1275 model, operated at a slit width of 1 urn. Argon was used as the internal163Figure 4.1 Schematic diagram of the apparatus used for the HG-AAS methodOPTICAL PATHof A.A. SpectrometerReaction CoilPeristaltic PumpSampleAcidNaBH4Gas-LiquidSeparatorDrainPressureRegulatorreported by Cullen and Dodd.5.2cm(I.D.)CSnH4ES0EEIt•)c.’JFGNoBH4QUARTZ 22cmxl .lmm(I.D.)HD4mm stopcockGRAPHITE FURNACETUBEWATER VACUUMFigure 4.2 Schematic diagram of the hydride generator used for HG-GFAAS.164purge gas. The tin hollow cathode lamp was purchased from Hamamatsu Photonicsof Japan. Analyses were carried out at the 224.6 nm spectral line.4.2.1.2 Batch hydride generation-graphite furnace atomic absorptionspectrophotometry (HG-GFAAS).The batch hydride generator used for the HG-GFAAS is a glass apparatusshown in Figure 4.2, and was modelled to be slightly different from the designreported by Sturgeon for arsenic and selenium determination, and later fortotal tin determination229. The lower portion of the batch hydride generator wasconstructed of a 25 mL Buchner funnel (Corning Glass Works, Corning, U.S.A) withmedium porous glass fit (pore size 10-15 J.Lm). This hydride generator was designedto accomodate larger volumes of reagents than the one reported by Sturgeon et a1,and the larger surface area of the glass fit should enable easier mixing of thereagents, and purging of the generated tin hydride out of the batch hydridegenerator. The tin hydride produced in the hydride generator was swept by anupward flow of nitrogen via a Teflon® tubing to a narrow quartz tube of innerdiameter 1.1 mm, which was inserted into the heated graphite tube of the graphitefurnace atomizer, aligned in the optical path of the atomic absorptionspectrophotometer. The graphite furnace tubes were pre-used Varian Techtron®graphite tubes, which were pre-coated with either sodium tungstate or palladiummodifiers. When no pre-used tubes were available, fresh graphite tubes whosepyrolytic coatings had been roughened by using an abrasive (sandpaper), were coated165with solutions of these modifiers, and used. Sturgeon have reported that preused graphite tubes are more efficient in trapping the tin hydride than fresh graphitetubes. The orifice on the wall of the graphite furnace tube was widened to a diameterof 2.3 mm to allow the insertion of the quartz tube. The graphite furnace atomizerwas a Varian GTA-95 instrument connected to a Varian 1275 atomic absorptionspectrophotometer. The spectral line and slit width used are as described in Section4.2.1.1.4.2.2 Materials and reagents.The following chemicals; sodium tungstate dihydrate (Analar grade), Lcysteine, sodium borohydride (Assured grade), potassium hydroxide (Aristar grade)hydrochloric acid (Analytical grade) were purchased from BDH Chemicals Ltd,Poole, England. Palladium powder was procured from Ventron Chemical Company,Danvers, Massachusetts, U.S.A. .Tin metal was obtained from Mallinckrodt ChemicalWorks, St Louis, Missouri, U.S.A.. Tort 1 (lobster hepatopancreas) standardreference material was obtained from the National Research Council, Canada.Hydrofluoric acid (doubly distilled in quartz) was obtained from Sea Star Chemicals,Victoria, Canada.1664.2.3 Methodology for the hydride generation atomic absorption spectrophotometry.4.2.3.1 Continuous hydride generation method (HG-AAS).The reagents were pumped by means of a peristalic pump into the reactioncoil (A in Figure 4.1), where mixing of the reagents and the production of the tinhydride occurred. A flow of nitrogen gas ensured the purging of the generated tinhydride into the gas-liquid separator (B in Figure 4.1) which was further purged bynitrogen gas. The tin hydride was swept into the heated T-shaped quartz cell whereatomization occurred. The absorbance reading was recorded after it became stable.All analyses by the HG-AAS method were carried out in triplicate.4.2.3.2 Batch hydride generation-graphite furnace method (HG-GFAAS).The batch hydride generator and the hydride transfer lines were silanized byusing a 10% (vlv) triethylsilane solution in toluene as follows:- with all the transferlines connected, and all taps closed except stopcock H at the bottom of the hydridegenerator, the quartz tube G was immersed in a solution of 10% triethylsilane intoluene with the water vacuum turned on. The triethylsilane solution was drawn intothe hydride generator, and the spent solution was then pumped out via stopcock FLThe hydride generator was then dried with a gentle flow of nitrogen, admittedthrough tap D, over a period of about fifteen minutes. This procedure minimized theadsorption of the tin hydride on the walls of the hydride generator.During the analysis, measured amounts of hydrochloric acid and the sample167were each pipetted onto the porous glass fit via a B24 joint at the top of the hydridegenerator, while an upward flow of nitrogen was maintained through tap DTable 4.1 Graphite furnace atomization program for tin determination by(HG-GFAAS).Step Temperature °C Time (s) Gas flow (Llmin)1 700 19 3.02 700 40 3.03 700 40 3.04 700 40 3.05 700 40 3.06 700 4.0 3.07 2700 4.0 0.0*8 2700 2.0 0.0*9 2700 1.0 3.0*= When absorbance measurement was taken.Steps 1-6 represent trapping and drying conditions.Steps 7-8 represent atomization conditions.Step 9 is clean up.168(Figure 4.2). Measured amounts of sodium borohydride solution were delivered intothe hydride generator by means of a peristaltic pump via tap F. The tin hydrideproduced in the hydride generator was swept by an upward flow of nitrogen admittedthrough tap D, to a narrow quartz tube of inner diameter 1.1 mm, which was insertedinto the heated graphite tube of the graphite furnace atomizer, and trapped by usingthe furnace program shown in Table 4.1.A gentle flow of argon maintained an inert atmosphere inside the graphitefurnace tube, except during the atomization step when the argon flow was stopped.After the tin hydride had been trapped in the graphite furnace tube, the nitrogenflow into the quartz tube was stopped, and the quartz tube was manually removedfrom the graphite furnace tube which was then quickly heated to 2700 °C, to atomizethe analyte. After each determination, the solution remaining in the hydridegenerator was pumped out, by using the water vacuum.4.2.4 Preparation of matrix modifiers and standard tin solutions.4.2.4.1 Preparation of palladium modifier.The palladium modifier solutions (2-10 % w/v) used to treat the graphitetubes were prepared by dissolving palladium metal in 1 mL of a warm mixture ofconcentrated hydrochloric acid and nitric acid (1:5 vlv), and diluting with 2 %ascorbic acid solution in a 5 mL volumetric flask.1694.2.4.2 Preparation of sodium tungstate modifier.Solutions of the sodium tungstate modifier (2 - 10 % w/v) were prepared bydissolving sodium tungstate dihydrate in de-ionized water.4.2.4.3 Preparation of standard tin solutions.Stock standard solutions were typically prepared by dissolving tin metal shotin 2 mL of a warm mixture of concentrated HC1 and HNO3 (1:1), and then dilutingthe resulting solution to 50 mL in a volumetric flask. Working standard solutionswere prepared by diluting appropriate amounts of the stock solution in 0.5 Maqueous HC1 solution. The working standard solutions used for quantitation wereprepared in 0.5 M HC1 solutions containing 2% L-cysteine.4.2.5 Optimum concentration of reagents used in the continuous hydride generationmethod (HG-AAS).The generation of tin hydride is pH dependent113. The optimum pH andreagent concentrations were established as follows:-various concentrations of sodiumborohydride in the range 0.5 - 2.5 % (w/v) were prepared in aqueous potassiumhydroxide solution (0.2% wlv). A standard tin solution in 0.5 M HC1, and differentconcentrations of hydrochloric acid in the range 0.1 - 1.0 M were each prepared indifferent volumetric flasks. When required, these reagents were pumped into thereaction coil of the continuous hydride generator by using the parameters shown inTable 4.2.170At a fixed concentration of hydrochloric acid, standardvarying concentrations of sodium borohydride, simultaneouslyreaction coil, the absorbance of the tin hydride produced wasatomic absorption spectrophotometer. The absorbance measuredindication of the yield of tin hydride. The reagent concentrationsabsorbance of tin hydride were then used for the determinationtin solution, andpumped into themeasured by thewas taken to be angiving the highestof total tin.Table 4.2 Operating conditions for the continuous hydrideabsorption specirophotometry (HG-AAS).generation atomicFlow rate SampleFlow rate HCLFlow rate NaBH4Purging gas flow rate4.4 mL/min4.4 mL/min4.4 mL/min0.6 L/min4.2.6 Use of L-cysteine to remove interferences.During the atomic absorption analyses of the marine animal samples, it wasobserved that the absorbance signal started to decrease as the analysis progressed,until it finally disappeared. This phenomenon was more noticeable when digestedenvironmental samples were introduced either into the batch or continuous hydridegenerators. Such behavior had previously been encountered by other workersincluding Brindle and Le236, Beach and Shradert13, Le Ii237, and Quevauviller171a!112. To eliminate this interference they added either L-cystine236 or L-cysteine113’7to the reaction mixture prior to hydride generation. Consequently, in the presentwork a study was carried out to find the optimum concentration of L-cysteine neededto prevent the disappearance of the tin absorbance.4.2.6.1 Optimum concentration of L-cysteine required to removeinterferences.Standard tin solutions (0.2 g/mL) containing 0.5- 3.0.ig/mL L-cysteine wereprepared in 0.5 M hydrochloric acid. The absorbance corresponding to the tinhydride produced from the reaction between the standard tin solutions and sodiumborohydride were measured by using HG-AAS.For the batch hydride generation method, no optimization was carried out, butthe use of the optimum concentration of L-cysteine obtained for the continuoushydride generator was sufficient to prevent the disappearance of the tin absorbance.4.2.7 Optimum conditions for the batch hydride generation-graphite furnaceatomic absorption specirophotometry (HG-GFAAS).The batch hydride generation method was optimized for concentration andvolume of reagents, trapping temperature, and trapping time of the tin hydride in thegraphite furnace tubes coated with 8% sodium tungstate modifier solution.1724.2.7.1 Optimization of reagent concentrations for HG-GFAAS.A standard tin solution (1 mL of 12 ng/mL tin solution), prepared in 0.5 Mhydrochloric acid, was added into the batch hydride generator, and then reacted withvarious volumes of 0.2 M hydrochloric acid solution, and 2 % (wlv) sodiumborohydride in 0.2 % potassium hydroxide solution. The absorbance of the generatedtin hydride was measured, and taken to be an indication of the yield of tin hydride.The results obtained for this optimization are discussed in Section 4.4.4.1.4.2.7.2 Optimization of trapping temperatures and trapping time for tinhydride in the graphite furnace tube.With the optimum volumes of reagents established, the trapping temperaturefor the generated tin hydride in the graphite furnace was varied, and the atomicabsorbance of tin hydride measured. The experiment was repeated at other trappingtemperatures.After the optimum trapping temperature had been established, the effect ofthe trapping time on the absorbance was also studied, by varying the trapping timeof the tin hydride at a constant trapping temperature and reagent concentrations. Theresults obtained in this study are discussed in Section 4.4.4.3 and 4.4.4.4.4.2.8 Treated graphite furnace tubes:- coating the graphite furnace tubes withsolutions of sodium tungstate and palladium modffiers.The method used in this study for the treatment of graphite furnace tubes with173matrix modifiers, is similar to the procedure described by Fritzsche Lai8.Pre-usedgraphite furnace tubes were soaked for 26 hours in aqueous sodium tungstatesolution (2 -10% wlv) or in a solution of palladium (2 - 10% w/v) in 2 % aqueouscitric acid. The preparation of the palladium and sodium tungstate modifiers isdescribed in Section 4.2.5. The soaked graphite furnace tubes were dried in an ovenat 125 - 129 °C for 4.5 hours. Prior to use, they were cleaned once by raising thetemperature of the graphite furnace to 3000 °C, and then conditioned by running thegraphite furnace program (shown in Table 4.1) four consecutive times.4.2.8.1 Optimum modifier treatment of graphite furnace tubes.A standard tin solution was used to produce tin hydride which was trapped inthe graphite furnace tubes treated with varying concentrations of sodium tungstate(2 - 10 % w/v) or palladium modifier (2 - 10 % w/v) solutions. The preparation ofthe sodium tungstate and palladium modifiers is described in Section 4.2.4.A plot of absorbance versus modifier concentration (Section 4.4.4.5) revealedthe optimum modifier concentration required to coat the graphite furnace tubes.4.2.8.2 Calibration curves for the HG-GFAAS method.Tin standards (2 - 14 ng/mL) in 0.5 M HC1 containing L-cysteine (2% w/v)were introduced into the batch hydride generator, and reacted with 0.2 M HCI (5mL), and 4 mL of 2 % sodium borohydride solution containing 0.2 % KOH, toproduce tin hydride which was trapped on sodium tungstate-treated graphite tubes.174The measured absorbances were piotted as a function of the concentrations of thestandard tin solutions, to obtain a calibration curve.4.3 Sample digestion and preparation.Freeze dried oysters or the standard reference lobster hepatopancrease, Tort1 (about 2.00 g) and 2% aqueous potassium hydroxide (20 mL) were placed in a 500mL round bottom flask fitted with an air cooled reflux condenser previouslydescribed by Dodd239 (Appendix D), and refluxed for lh 45 mm. The contents of theround bottom flask were cooled, and concentrated sulphuric acid (4 mL of 12 M) andconcentrated nitric acid 30 mL of 15 M) were added to the round bottom flask, andfurther refluxed until all solution had gone into the reflux condenser, and the residuein the round bottom flask has charred. Heating was stopped, and after a few minutes,when the solution in the reflux condenser had dripped back into the round bottomflask, refluxing was resumed until the solution became clear and colorless, or verylight yellow. The round bottom flask was cooled, and de-ionized water(10-20 mL) wasadded through the reflux condenser. Reflux was then continued until the solutionturned colorless. The solution was cooled and then transferred to a 250 mL glassbeaker where the solution was evaporated down to about 10 mL, by using a hotplate. The solution was transferred to another 250 mL beaker made of Nalgene®,followed by the addition of 1 mL hydrofluoric acid. The solution was furtherevaporated on the hot plate to about 5 mL. Concentrated hydrochloric acid, and deionized water (50 mL) were added to the Nalgene® beaker, and further heated until175the volume of the solution has reduced to about 30 mL. The solution was cooled,transferred to a 50 mL volumetric flask containing L-cysteine, and made up to themark with 0.5 M hydrochloric acid solution to form the digested sample in 2% w/vL-cysteine solution.A blank solution containing all the reagents used for sample digestion was alsodigested, by following the same digestion procedure described for the sample.4.4 RESULTS AND DISCUSSION.4.4.1 Optimum concentrations of sodium borohydride and hydrochloric acidnecessary for the production of slnnnane in the continuous hydride generator.The effect of the concentrations of hydrochloric acid and sodium borohydrideon the generation of tin hydride, as monitored by measuring the absorbance of thegenerated hydride, is shown in Figure 4.3. The error bars on all the graphs in thisChapter are the standard errors for three replicate determinations. At all the sodiumborohydride concentrations studied, more SnH4 was generated as the sodiumborohydride concentration was increased. At 2% sodium borohydride concentration,a maximum is reached, and further increase in sodium borohydride concentrationleads to decreased SnH4 production as shown by a decrease in absorbance at 2.5 %sodium borohydride. As the hydrochloric acid concentration is increased, theabsorbance of the generated tin hydride decreased. L-cysteine was not used in thisoptimization study.1768IFigure 4.3 Effect of sodium borohydride and HC1 on the absorbance of tm hydrideI I0.4 -0.3 -0.2-0.1 -0,0 --I-0.2 0.4 0.6 0.8Concentration of HC1 (M)1 .00.0A0.5•1.502.01 .2% sodium borohydride% sodium borohydride% sodium borohydride% sodium borohydridev 2.5 % sodium borohydrideproduced from 4 tg/mL tin solution.1774.4.2 Optimum concentration of L.cysteine required to eliminate interfereDces inHG-AAS.During the HG-AAS analysis of the marine animal samples, the tinabsorbance started to decrease as the analysis progressed, especially when the oysteror Tort 1 digested solutions were introduced either into the batch or the continuoushydride generators or when the quartz cell of the continuous hydride generatorbecame dirty with an insoluble material.Interferences capable of causing the decrease of the tin absorbance signals canbe encountered in two stages of the hydride generation-atomic absorption analysis:(a) In the hydride generator, where other metal ions could compete with Sn forborohydride.(b) In the heated quartz cell, where the formation of refractory tin carbide, whichdoes not atomize at the temperature of the air-acetylene flame would reduce tinabsorption.Brindle and Le6, Le et a17, Nakahara24° and Thompson et al24’ havereported that transition metal ions such as Fe(II), Fe(III), Co(II), Ni(II), and Cu(II)cause serious reduction of tin absorbance signals. Such interferences that inhibit theformation of SnH4 had previously been eliminated by using L-cysteine113’7or Lcystine236. According to these authors113’7,L-cysteine also decreased the pHdependency of the tin hydride formation.Therefore, a study was carried out to find the optimum concentration of Lcysteine required to improve tin absorbance.178The optimum concentration of L-cysteine was found by analyzing a 0.2 .ig/mLtin standard solution containing varying concentrations of L-cysteine, and plotting theabsorbances against L-cysteine concentrations (Figure 4.4). Figure 4.4 indicates thatthe optimum concentration of L-cysteine is about 2 % (wlv). The optimumconcentration of L-cysteine is higher than the concentration reported by Beach andShrader113 and Le tai237 in their methodologies, to improve tin absorbance. Bothauthors used a 1 % L-cysteine solution.Consequently, all standard tin solutions and digested animal samples weredissolved in solutions containing 2% L-cysteine.0.200.150.100.050.003.50.0 0.5 1 .0 1 .5 2.0 2.5 3.0Concentration of L-cysteine (% wlv)Effect of L-cysteine on absorbance of tin hydride.Figure 4.41794.4.3 HG—AAS determination of total tin in oysters and standard reference material(Tort 1).Digested sample solutions in 2% L-cysteine solution were pumped into thereaction coil of the continuous hydride generator, where a reaction occured betweensodium borohydride and tin, to produce the stannane which was detected by theatomic absorption spectrophotometer. Quantitation of the total tin content of thesamples was accomplished by using the standard addition method as follows; 5 mLof the digested sample in 2% L-cysteine solution were spiked into 10 mL of standardtin solutions (0 - 8 tg/mL Sn) containing 2% L-cysteine in 0.5 M HC1 solution. Thedigested blank solution (5 mL) was also spiked into another set of calibrationstandards (0 - 8 jg/mL Sn) containing 2% L-cysteine in 0.5 M HC1. Each mixtureof the digested sample solution and the tin standard solution (4 mL) was pumpedinto the reaction coil of the hydride generator, and the absorbance of the generatedtin hydride was measured by the atomic absorption spectrophotometer. Similarly, theabsorbance of any tin hydride produced from each mixture of the digested blank andthe standard tin solutions was also measured. Two replicate determinations of eachsample mixture were carried out. The total tin contents of the samples were thencalculated, after the blank values had been subtracted.Quantitation by using the more difficult and time consuming standard additionmethod was preferred in this study, because repeated analyses of the certifiedreference material Tort 1, by the normal calibration method consistentlyoverestimated its total tin content by about three fold. This situation could not be180improved upon, neither by the use of background correction nor blank subtraction.Without the addition of 2% L-cysteine into the sample and standard solutions,quantitation would not be possible, because the tin absorbance was completelysuppressed in some determinations. The mechanism of action of L-cysteine is notknown with certainty.The total tin content of the samples and the standard reference materials areshown in Table 4.3.The total tin content of the standard reference material Tort 1 obtained inthis study is in the range previously reported by Sturgeon et al229. Therefore, thedigestion method and the HG-AAS method employed in this study are suitable forthe determination of total tin in marine animals.Since the normal calibration method of quantitation gave a much higher totaltin value than the certified value for Tort 1, another atomic absorption methodcapable of reproducing the total tin content of Tort 1, by normal calibrationprocedure was sought, because of the rapidity and ease of this quantitation method.Two non-conventional hydride generation-atomic absorption methods; acontinuous HG-AAS method developed by Le et aI7 and a batch HG-GFAASmethod reported by Sturgeon a.i229,were considered. The batch HG-GFAAS waspreferred over the non-conventional continuous HG-AAS method of Le et al7because, their method was not validated for the quantitative determination of tin, asneither its reproducibility nor detection limit was reported. Also, the method7 is notcapable of reaching the very high atomization temperatures characteristic of the181Table 4.3 Total tin content of samples analyzed by the HG-AAS method.Sample Origin Total tin content Certified(g/g dry wt)a value (ig/g dry wt)Tort 1b NRC, Canada 0.16 ± 0.04 0.139 ± 0.011c0.144 ± 0016dPacific oyster Cambell River 0.34 ± 0.09Crassostrea gigasPacific oyster Fanny Bay 0.35 ± 0.03Crassostrea gigasPacific oyster Jervis Inlet 0.13 ± 0.01Crassostrea gigasa=Total tin content and the standard deviation for 3 replicate determinationsb =Lobster hepatopancrease, a standard reference material from the NationalResearch Council, Canada (NRC).c=Value certified by NRC, Canada.d=Value reported by Sturgeon Lal229.batch HG-GFAAS method. Therefore, molecular absorption or non-atomization ofrefractory tin compounds in the air-acetylene flame, in the quartz furnace of the non-conventional continuous HG-AAS might pose a problem.182Consequently, a batch HG-GFAAS apparatus (Figure 4.2) modelled on theprinciple reported by Sturgeon but slightly different in design was constructedand optimized for total tin determination.4.4.4 Batch hydride generation-graphite furnace atomic absorptionspecirophotometry (HG-GFAAS).The use of the continuous HG-AAS method, has a major disadvantage ofconsuming large amounts of samples and reagents, and is therefore wasteful andexpensive. Conversely, the batch hydride generation method consumes very smallamounts of samples and reagents. The small amounts of tin hydride generated aresuitable for trapping on a graphite furnace tube, where it is preconcentrated prior toatomization. The preconcentration step, and the ability of the inside surface of thegraphite tube to reduce some refractory compounds, are expected to increase thesensitivity and the detection limit of this method.For normal operation in the graphite furnace mode, the steps involved are:sample drying, ashing, atomization, and tube cleaning. During the drying stage,solvent or water is removed from the sample. At the ashing step, organic andinorganic matrices are removed. However, in the hydride generation-graphite furnacemethod (HG-GFAAS), organic and inorganic matrices are minimized. At theatomization step, free atoms of the analyte are generated in the graphite tube, andtheir absorbances are measured by the atomic absorption spectrometer. During thegraphite furnace operation, the incandescent graphite tube is protected from183excessive corrosion by an upward flow of an inert gas such as argon or nitrogen(internal purge).The use of the graphite furnace tube to trap or adsorb tin hydride has beendemonstrated by Sturgeon Lili229’5and was the basis for a HG-GFAAS methodreported by these authors229’5.The extension of this methodology to trap tinhydride in graphite furnace tubes precoated with sodium tungstate, and palladiummodifiers is described in this section. In the method reported by Sturgeon Li229,no modifiers were used, probably because there was no interference from theirsample matrix. In this study, the use of perchloric acid for sample digestion as usedby Sturgeon et a1229 was avoided because of its explosive nature, instead KOH,HNO3,H2S04,and HF were used in various stages of the sample digestion (Section4.3). The difference in reagents used for sample digestion may have contributed tothe extent of interferences observed during the HG-GFAAS analysis reported in thepresent study. The initial approach taken to remove these interferences involved themanual injection of solutions of sodium tungstate or palladium modifiers into thegraphite furnace tube prior to every absorbance measurement. 1_ater, the use ofgraphite furnace tubes pre-treated with solutions of palladium or sodium tungstate,and the presence of L-cysteine in the batch hydride generator, made it possible toanalyze the environmental samples. The use of graphite furnace tubes pre-coatedwith solutions of these modifiers eliminates the inconvenience of manually injectingmodifiers into the graphite furnace tube during each analysis.1844.4.4.1 Optimum concentrations of reagents needed for tin hydrideproduction in the HG-GFAAS method.At a trapping temperature of 700°C, standard tin solution in 0.5M HC1 (1 mLof 12 ng/mL Sn), and measured amounts of 0.2 M hydrochloric acid (2 - 20 mL)were pipetted into the batch hydride generator via a B24 joint at the top of thehydride generator. Measured amount of aqueous sodium borohydride (4mL of 2 %w/v in 0.2% KOH solution) was pumped into the batch hydride generator to reactwith a standard tin solution. The absorbance measurements represent the amount oftin hydride produced (Table 4.4).Table 4.4 Reagent ratios needed to maximize tin hydride generationVolume 0.2 M HC1 (mL) Volume 2 % NaBH4 (mL) Absorbance (±)a2 4 0.038 ± 0.0075 4 0.040 ± 0.00310 4 0.040 ± 0.00720 4 0.028 ± 0.001a=Standard error for three determinations.All the volume ratios of the reagents examined gave about the same absorbancevalues when 2 - 10 mL of 0.2 M HC1, and 4 mL sodium borohydride were used for0.040.030.020.01185tin hydride production (Table 4.4). A probable reason for this observation is that inall the cases, the sodium borohydride was present in excess, therefore the reactionwent to completion at all the reagent ratios studied. Since the reagent ratios used didnot appear to be critical for tin hydride production, provided the sodium borohydridewas in excess, all batch hydride generation, experiments were carried out at thereagent ratio of 0.2 M HC1 (5 mL) : 2% sodium borohydride (4 mL).4.4.4.2 Optimum flow rate of sodium borohydride into the batch hydridegenerator.The effect of the flow rate of sodium borohydride solution into the batchhydride generator, on the absorbance of tin hydride is shown in Figure 4.5. As the0.054.0 4.5 5.0 5.5 6.0 6.5 7.0NaBH4 flow rate (mLlmin)Figure 4.5 Effect of sodium borohydride flow rate on absorbance. (The width ofthe bars is arbitrary. The error bars are std error for 3 determinations).186sodium borohydride flow rate was increased from 4.4 to 6.3 mL/min, a maximumabsorbance was observed at about 5.40 mL/min. This indicates that the flow rate ofthe sodium borohydride into the hydride generator affects the production of the tinhydride. In their work, Sturgeon 229 used a flow rate of 4 mL/min to deliver 2mL of sodium borohydride solution into their hydride generator. The difference inflow rate between the batch hydride generator used in this study and the onereported by Sturgeon et a!229 may be due to the difference in the size of the twohydride generators.4.4.4.3 Optimum temperature for trapping tin hydride in the pre—treatedgraphite furnace tubes.The effect of temperature, on the ability of the graphite furnace tubes to traptin hydride was studied by measuring the absorbance of tin hydride produced froma reaction between standard tin solution (1 mL of 14 g/mL solution) and sodiumborohydride (4 mL of 2% solution), as the temperature of the graphite furnace tubeis varied. The result obtained is shown in Figure 4.6.The absorbance of the tin hydride, as the temperature of the graphite furnacetube is varied, is an indication of the trapping efficiency of the graphite furnace tube.In the temperature range studied, maximum trapping efficiency was obtained atabout 700 °C. This trapping temperaturç is lower than the value reported bySturgeon et a!229, by 100 °C. The lower trapping temperature established in thepresent study may be due to the pre-treatment of the graphite tubes with sodium187tungstate matrix modifier, and is expected to prolong the “life span” of the graphitetube.0.080.07 -Ie 0.06 -0,05 -0.04--100 200 300 400 500 600 700 800Trapping temperature °CFigure 4.6 Effect of trapping temperature on the atomic absorbance of tin hydride.4.4.4.4 Optimum trapping time.A study was carried out to find the trapping time needed to produce maximumabsorbance. Figure 4.7 shows the effect of trapping time on the absorbance of tinhydride. Trapping efficiency as monitored by absorbance measurements wasmaximum at 250-260 seconds. Thereafter, the trapping efficiency decreased. TheI I I I188decrease in absorbance as trapping time increased beyond 260 seconds may be dueto the desorption and escape of the tin hydride from the graphite tube.0.05 -0.04 -0.03 -0,02-—260 270 280Figure 4.74.4.4.5 Pre-treatment of graphite furnace tubes with modifiers.Vickrey et a1242 have coated some carbide forming elements such aszirconium, chromium, and molybdenum onto graphite furnace tubes by soaking thegraphite furnace tubes in solutions of these elements. This coating method242 waseffective in reducing atomization interferences in the GFAAS determination oforganotin compounds. Another method of coating graphite furnace tubes withcarbide forming elements has been described by Almeida and Seitz243.This methodinvolved soaking the graphite furnace tubes in solutions of titanium, molybdenum,I I I t I I II I200 210 220 230TrappingEffect of trapping time240 250time (s)on absorbance.189or tungsten salts under reduced pressure, in a vacuum line. This method wasdescribed by the authors as being more efficient in causing the penetration of thegraphite furnace tubes by the modifier, than the method of Vickrey et a1242.According to Almeida and Seitz243, the modifiers tungsten and titanium coat on thegraphite furnace tubes as oxides, but are converted to the carbides at the highgraphite furnace temperatures.The coating of graphite furnace tubes with carbide forming elements has alsobeen described by Lagas244.His method involved the injection of an aqueous solutionof lanthanum chloride into a graphite furnace tube. Drying, ashing, and atomizationprograms of the graphite furnace converted the injected lanthanum salt to its carbide.The physical basis for the action of carbide forming elements in GFAAS hasbeen described by Lagas244. According to this author, the carbide coating preventsphysical contact between the graphite tube and the analyte thereby preventingcarbide formation by the analyte.Although the use of palladium modifiers to coat GFAAS tubes has not beenreported, it was thought feasible to explore such methodology. It is expected that themechanism of modifier action would be the same as when palladium solution ispremixed with the analyte as is normally done in GFAAS. Its use as a modifier in theanalyses of an increasing number of elements has been reported245’678Palladium does not act by forming carbides. According to Volynsky249,palladium acts by forming intermetallic compounds and solid solutions with theanalyte in the graphite furnace. To function as a modifier, palladium salts are usually190reduced to Pd(0) by using either a solution of citric acid or ascorbic acid.In the present study, palladium and sodium tungstate modifiers were coatedonto graphite furnace tubes, by soaking the graphite furnace tubes in solutions ofthese modifiers over a period of 26 hours as described in Section 4.2.8The effect of the concentration of the modifiers on the absorbance of tinhydride, produced by reacting 1 mL of 14 ig/mL tin solution with 4 mL of 2%sodium borohydride in the batch hydride generator is shown in Figures 4.8 and 4.9.For the graphite furnace tubes coated with sodium tungstate, treatment with8 % sodium tungstate gave maximum absorbance, while treatment with 4% palladiummodifier gave maximum absorbance for the palladium-treated graphite tubes.The exact amount of the modifier coated on each graphite furnace tube wasnot determined, but it was assumed that the graphite furnace tubes would absorbsimilar volumes of the modifiers, hence the amount of modifier coated onto thesetubes should be proportional to the concentration of the modifier solution in whichthe graphite furnace tubes were soaked. This assumption seems reasonable becausethe pre-used graphite furnace tubes were in about the same physical condition, andwere from the same batch supplied by the same manufacturer.Comparatively, at the optimum modifier concentrations, sodium tungstatetreated graphite furnace tubes gave better tin absorbance than palladium treatedtubes (Table 4.5). Also, analyses with the palladium treated graphite tubes wascharacterized by a loud popping sound resulting from the ignition of the air-hydrogenmixture in the heated graphite tube. Therefore, the graphite tubes treated with 8%I I I I0 2 4 6 8 10 12Concentration of sodium tungsiate (% wlv)Figure 4.8 Effect of sodium tungstate concentration on absorbance.191I I—- I II0.070.06 -0.05-0.04I I I I I0.06 -0.05 -0.040.030.020.01 -0.00 I I I I0 2 4 6 8 10 12Concentration of palladium (% w/v)Effect of palladium on absorbance.Figure 4.9192sodium tungstate were preferred over those treated with the 4% palladium solution.Table 4.5 Comparison of palladium and sodium tungstate-lreated graphitefurnace tubes showing the atomic absorbance of tin hydride generatedfrom 14 JLg/mL tin solution (1 mL).Graphite furnace tube treatment Absorbance (±)a4% Palladium treatment 0.054 ± 0.0028% Sodium tungstate treatment 0.062 ± 0.003a=Standard error for three determinations.4.4.4.6 Determination of total tin content of a standard reference material bythe HG-GFAAS method.About 2.00 g of Tort 1 (lobster hepatopancrease), a standard referencematerial was digested as described in Section 4.3. A blank solution of all thereagents used to digest Tort 1 was similarly digested. The digested Tort 1 solutionwas transferred to a 25 mL volumetric flask containing L-cysteine, and dissolved inde-ionized water, so that the concentration of L-cysteine was 2% w/v. A solution ofthe digested blank, in 2% L-cysteine was similarly prepared. Aliquots of the digestedsample or the digested blank solutions (3 mL), sodium borohydride (4 mL) and HC1(5 mL) were reacted in the batch hydride generator, and the absorbance of the193generated tin hydride was measured.Quantitation of total tin in Tort 1 was carried out by the normal calibrationmethod, by using the calibration curves obtained as described in Section 4.2.8.2.The total tin content obtained by using HG-GFAAS is shown in Table 4.6.Table 4.6 Total tin content of a standard reference material Tort 1 obtained bydifferent authors.Technique (Source) ValueHG-GFAAS, (the present study) 0.102 ± 0.018aHG-GFAAS, (Sturgeon ii232) 0.144 ± 0.016k’HG-GFAAS, Isotope dilution ICP-MS 0.139 ± 0.01 1b(NRC, Canada)a= Standard deviation of two replicate determinations.b = Standard deviation, but number of replicate determinations was not givenby the authors.The total tin content of Tort 1 determined by the batch HG-GFAAS methoddescribed in this study lies close to the lower range of values certified for Tort 1(Table 4.6). The performance of this batch hydride generator may be improvedfurther, by using other optimization techniques, such as the simplex method.194However, the batch HG-GFAAS method described in the present study offers analternative to the method described by Sturgeon et a1229, especially in situationswhere matrix interferences are a problem. This is because their method229 wasreported to operate without encountering any interferences, therefore its operationin situations such as in the present study where intereferences posed a problem, hasnot been tested.When the continuous HG-AAS method and the batch HG-GFAAS used inthis study are compared, the latter offers a faster method of quantitation by normalcalibration. The continuous HG-AAS method is time consuming, involving standardadditions. Other comparative data on the two atomic absorption methods of analysisused in the present study are shown in Table 4.7.Table 4.7 Comparison of figures of merit obtained with the two atomicabsorption spectrophotomelric methods used in this study.HG-AAS HG-GFAASDetection limit 7.5 ng/ mL 1.8 ng/mLPrecision (for 10 runs) 4.3% 3.3%195CHAPTER 5SUMMARY AND CONCLUSIONS.Studies involving the analyses of organotin compounds in marine organismsof British Columbia, Canada, and the effect of organotin compounds on thepermeability of model biological membranes have been presented in this thesis.Analysis of organotin compounds in marine organisms by GC-MS SIM affordsa very specific technique for the identification and quantitation of these organotincompounds by using the peculiar isotope pattern for tin compounds. Thismethodology is therefore able to distinguish organotin compounds from othercompounds that may co-elute with them from the GC.The major organotin pollutants found in this study for the coastal areas ofBritish Columbia were tributyltin and dibutyltin species. Dicyclohexyltin levels of 3.5ng/g (wet wt as Sn) and 21.3 ng/g (wet wt as Sn) were found in only two locations,namely Wreck Beach, Vancouver and Anyox. Therefore, pollution fromdicyclohexyltin species is not widespread in the coastal areas studied.Contamination of mussels and clams by tributyltin and dibutyltin species iswidespread, although some locations such as Hastings Arm, Hilton Point Kitimat,Alice Arm and Holberg Sound show no organotin pollution. The butyltin bodyburden for Blue mussels in the contaminated areas sampled range from 14.4 ng/g to37.3 ng/g (wet wt as Sn) for tributyltin species and 6.7 to 67.3 ng/g (wet wt as Sn)for dibutyltin species. The trend observed is that most Blue mussels showed the196presence of both dibutyltin and tributyltin. This is expected because dibutyltin speciesare degradation products of the tributyltin species, and should be present if tributyltincompounds are present. The Soft shell clams analyzed showed tributyltin levelsranging from 0.7 to 19.4 nglg (wet wt as Sn). The dibutyltin species were mostly notdetected even in the Soft shell clams that were contaminated by tributyltin species.This observation is contrary to expectation, and may indicate that dibutyltin speciesare easily eliminated from the soft shell clams when compared to the Blue mussels.For the other species of mussels and clams analyzed, paucity of samples could notallow for any trend to be established in terms of preferential accumulation oftributyltin or dibutyltin species.The study of tributyltin body content of Soft shell clams from Quatsino Soundshowed a significant reduction in levels from 26.3 ng/g (wet wt as Sn) to 5.5 nglg(wet wt as Sn) over a period of three years. This reduction in the concentration oftributyltin species may be due to the Canadian Government’s regulation of the useof tributyltin compounds, which came into effect in 1989.When compared to some other locations of the world contaminated bytributyltin species, the mussels analyzed in this study show comparable pollution tomussels from San Diego Bay, U.S.A.122and Tokyo Bay, Japan”’4.The use of 1H NMR spectroscopy to study the effect of organotin compoundson the permeability properties of model biological membranes was presented inChapter 3 of this thesis. This represents the first application of proton 1H NMR tostudy the permeability properties of butyltin-EPC liposomes. Direct study of the197permeation of the organotin compounds from the model biological membranes wasmade difficult by their high hydrophobicity and low aqueous solubility. The use ofdimethylarsinic acid (DMA) as a permeability probe afforded information on howthe organotin compounds affect the permeability of model biomembranes. Whentributyltin chloride was added to the extraliposomal compartment of an EPCliposome, the permeability of the liposomes to dimethylarsinic acid greatly increased.The mechanism of the increased permeability could not be determined because thediffusion equations [3.0] and [3.26] (Chapter 3, section 3.9), developed to describethis efflux behavior were unable to distinguish between situations in which there is100% passive diffusion and 100% facilitated diffusion. However, the increasedliposome permeability shows a linear relationship with the concentration oftributyltin chloride present in the extraliposomal compartment according to therelationship Y=8.01 x 104X + 1.46 x i(I, where Y is the efflux rate constant andX is the concentration of tributyltin chloride in the extraliposomal compartment.The permeability coefficient for DMAH efflux from EPC liposomesdecreased from 1.7 x 10-8 cm/s to 4 x i0 cm/s. when monobutyltin trichloride wasadded to the extraliposomal compartment of EPC liposomes. The decreasedpermeability of the EPC liposomes to DMAH in the presence of monobutyltintrichloride does not show a linear relationship. Beyond 16.7 jM monobutyltintrichloride added into the extraliposomal compartment, no further decrease inpermeability coefficient was observed.When trimethyltin hydroxide was added to the extraliposomal compartment198of EPC liposomes, the efflux of DMAH was by passive diffusion while the efflux ofDMK (another species of DMA present in solution at pH 7.4) was by facilitateddiffusion mediated by trimethyltin cation. The rate constant for the passive diffusionof DMAH (2.5 x iO Is) when in contact with trimethyl hydroxide was lower thanin the absence of trimethyltin hydroxide (1.1 xlO2 Is).For the tributyltin chloride-EPC liposomes (TBT-EPC), it was found thatDMAH permeated by passive diffusion, while DMK permeated by facilitateddiffusion if tributyltin chloride was absent in the extraliposomal compartment. Thisfacilitated diffusion was mediated by the tributyltin cations. The rate constant for thepassive diffusion of DMAH was dependent on the tributyltin chloride content of theliposomes, and varied from 3.2 x i(i Is to 1.7 x i0 Is for TBT-EPC A and TBTEPC B liposomes of composition 5 g TBT 0.2 g EPC , and 1.5 jg TBT 0.2 gEPC respectively.The rate of facilitated diffision of DMK encountered in this study, was foundto be controlled by the ratio of the formation constant to the dissociation constant(B/), of the trialkyltin-DMA complex. The ratio BI was found to be 2.5, and 2.1for the trimethyltin-DMA and tributyltin-DMA complexes respectively. Whentributyltin chloride was present in the extraliposomal compartment of a TBT-EPCliposome, the ability of tributyltin cation to mediate facilitated diffusion was removed.The activation energy for the passive efflux of DMAH from the TBT-EPCliposomes was also concentration dependent and decreased as the tributyltin chloridecontent of the liposome increased. The activation energies obtained were 64.4199U/mo! and 52.3 kJ/mol for TBT-EPC A and TBT-EPC B liposomes respectively.The Arrhenius pre-exponential factor for the TBT-EPC liposomes did notshow much variation with the tributyltin chloride content of the liposome, and wascalculated to be 2.8 x 10-8 and 4.0 x 10-8 /s for TBT-EPC A and TBT-EPC Brespectively.For the monobutyltin trichloride-EPC liposomes (MBT-EPC), the efflux ofDMA was by passive diffusion irrespective of whether monobutyltin trichloride waspresent in the extraliposomal compartment or not. The permeability coefficient ofDMAH for the MBT-EPC liposomes showed slight dependence on the monobutyltintrichioride content of the liposomes, and was 4.9 x i0 cm/s and 4.4 x i0 cm/s forMBT-EPC A and MBT-EPC B liposomes of composition 0.5 g MBT: 0.2 g EPC,and 1.5 g MBT: 0.2g EPC respectively. The activation energy for the efflux ofDMAH is 106.8 and 121.5 kJ/mol for MBT-EPC A and MBT-EPC B respectively.This indicates that the MBT-EPC liposomes become less permeable with increasingconcentration of monobutyltin trichioride in the liposomes. The Arrhenius preexponential factors calculated for the these liposomes are 1.1 x 10-8 and 1.0 x 10-8/s for MBT-EPC A and MBT-EPC B liposomes respectively.From the results of these permeation experiments, it is concluded thattributyltin chloride and monobutyltin trichioride exert different effects on thepermeability of both EPC liposomes and butyltin chloride-EPC liposomes. Whereastributyltin cation has the ability to act as a carrier for DMK, the monobutyltin cationhas no such ability. The present study is the first report of the facilitated transport200of an environmentally important compound, such as dimethylarsinate by trialkyltincation.The observation that the Arrhenius pre-exponential factor for the butyltinchloride-EPC liposomes are approximately constant for each type of liposomes, butdifferent for the TBT-EPC and the MBT-EPC liposomes tends to support theconclusion that tributyltin and monobutyltin species exert different effects on thepermeability properties of membranes. This difference may contribute to thedifferent toxic effects observed for these butyltin compounds on marine organisms.Although 1H NMR spectroscopy is a useful technique for studying the effectof organotin compounds on model membranes, its low sensitivity precludes the directstudy of organotin permeation at very low concentrations. There is also the possibilitythat the spectroscopic shift reagent used to distinguish between the proton resonanceinside and outside the liposomes may also modify the properties of the liposomes,thereby influencing the results. The use of a radioisotope technique which wouldallow for low level organotin permeation studies would be desirable, to check on theresults obtained in this investigation.Chapter 4 of this thesis is concerned with the determination of total tin inoysters from some locations in British Columbia, Canada. Two types of hydridegeneration-atomic absorption methods were employed for total tin determination: acontinuous HG-AAS and a batch HG-GFAAS method. 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Volynsky, A., Analyst 116, 145 (1991).220APPENDiX AMAP OF BRiTISH COLUMBIA, CANADA, SHOWING SOME LOCATIONSSAMPLED FOR ORGANOTIN POLLUTION.50221APPENDiX BTHE NMR SPECTRAL ACQUISITION AND WATER SUPPRESSIONPARAMETERS FOR THE EFFLUX OF DIMETHYLARSINIC ACID FROMLIPOSOMES.;Baz:AU PROGRAM FOR DIFFUSION EXPERIMENTSII2ZEVD1DOHG00=1 DOWR#1IF #1IN=2EXITVD VDLIST.0011-27 appropriate delays (seconds)1 0.001 10 1116.8 19 1716.82 216.8 11 1116.8 20 3516.83 216.8 12 1116.8 21 3516.84 216.8 13 1116.8 22 3516.85 216.8 14 1716.8 23 3516.86 516.8 15 1716,8 24 3516.82227 516.8 16 1716.8 25 3516.88 516.8 17 1716.8 26 3516.89 516.8 18 716.81 27 3516.8VD; the variable delay was obtained by subtracting the time required to obtain 48scans from the intended delay. The time required for the NMR spectrometer toobtain 48 scans was 83.2 seconds.Other NMR parameters were:PW=3DP=OLDR INITIAL= 16 FINAL =8Dl =0.5D3=3OSPW=PO=3.OONS=48DE=77.50DS=2Dl is the duration of the presaturation pulseD3 is the pulse delay timeDS is the dummy scanPW is the pulse widthDR is the digitizer resolutionDP is the decoupler power223APPENDIX CMICHAELJS-MENTONS EQUATIONS FOR ENZYME KINETICSFor enzyme catalyzed reaction:k1 k2E+S”ES-’Product+Ek1where E is the enzyme and S is the substrate.[ES]- k1[E][S]k1 +[ES] is the concentration of complexed enzymewhere (k1 + k2)/k, = kmkm is Michaelis constant.The rate of the forward reaction is:- V[S]km + [SI224APPENDIX DWET ASifiNG APPARATUS WiTH AIR COOLED REFLUX CONDENSERUSED FOR DIGESTION OF MARINE ANIMALS.Wet ashing apparatus (Taken from reference 242) showing the following parts (A)Teflon cylinderical plugs, (B) Teflon diffusion funnel, (C) Teflon stopper withcapillary, (D) 500 mL round bottom flaski.d 15 mmAB

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