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Nelson, John Christian
Doctor of Philosophy - PhD
University of British Columbia
The biotransformations and the biomobility of monomethylarsonate (MMA) and dimethylarsinate (DMA) in the marine environment were investigated. The biotransformations were studied by exposing blue mussels (Mytilus edulis) to either [³H]-MMA or [³H]-DMA in seawater tanks. Metabolic products including arsenobetaine ((CH₃)₃As⁺CH₂COO⁻) were identified in both the seawater and the mussels by using chromatographic techniques. The HPLC columns were coupled to a scintillation counter which acted as the detector. Arsenobetaine was also produced in seawater tanks which did not contain mussels which indicates that arsenobetaine is biosynthesized by microscopic organisms which are naturally occurring in seawater. It is likely that the arsenobetaine found in mussels is not a metabolic product of the mussels. The biomobility ofDMA and MMA was studied by measuring the rate at which these arsenicals diffused through liposomes as models for biological membranes. (Previously studies have shown that both DMA and MMA enter cells via slow passive diffusion.) Traditional techniques employing both radiolabelled permeants and liposomal membranes showed that DMA was much more permeable to the membranes than MMA. A ¹H-NMR spectroscopic technique employing shift agents to differentiate the spectroscopic signals of the permeant on either side of the liposomal membrane, was developed to measure the rate of diffusion of molecules across membranes. This technique was applied to DMA and MMA. Permeability coefficients of(5.8 ± 1.0) x 10⁻⁸ cm s⁻¹ and (7.8 ± 2.7) x10⁻⁹ cm s⁻¹ at 24°C were measured for the neutral forms of DMA and MMA, respectively. In addition, the effects ofpH, temperature, and membrane composition on the diffusion coefficients for DMA and MMA through the liposomal membrane were investigated. It was determined that only the neutral form of these weak acids permeates across the membrane and that cholesterol decreases the rate of permeation of DMA.
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THE BIOTRANSFORMATIONS AND BIOMOBU.ITY OFMONOMETI[YLARSONATE AND DIMETHYLARSINATE IN THE MARINEENVIRONMENTbyJOHN CHRISTIAN NELSONB. Sc., University ofBritish Columbia, Canada, 1989A THESIS SUBMITTED IN PARTIAL FULFILLMENTOF THE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIESDEPARTMENT OF CHEMISTRYWe accept this thesis as conformingto the required standardTHE UNWERSITY OF BRITISH COLUMBIA1993(c) John Christian Nelson, 1993In 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 Ce\ 5[The University of British ColumbiaVancouver, CanadaDate 4J7DE-6 (2/88)11ABSTRACTThe biotransformations and the biomobility of monomethylarsonate (MMA) anddimethylarsinate (DMA) in the marine environment were investigated.The biotransformations were studied by exposing blue mussels (Mytilus edulis) toeither[3H1-MMA or[3H]-DMA in seawater tanks. Metabolic products includingarsenobetaine ((CH)A.sCH2COOjwere identified in both the seawater and themussels by using chromatographic techniques. The HPLC columns were coupled to ascintillation counter which acted as the detector. Arsenobetaine was also produced inseawater tanks which did not contain mussels which indicates that arsenobetaine isbiosynthesized by microscopic organisms which are naturally occurring in seawater. It islikely that the arsenobetaine found in mussels is not a metabolic product of the mussels.The biomobility of DMA and MMA was studied by measuring the rate at whichthese arsenicals diffused through liposomes as models for biological membranes.(Previously studies have shown that both DMA and MMA enter cells via slow passivediffusion.) Traditional techniques employing both radiolabelled permeants and liposomalmembranes showed that DMA was much more permeable to the membranes than MMA.A ‘H-NMR spectroscopic technique employing shift agents to differentiate thespectroscopic signals of the permeant on either side of the liposomal membrane, wasdeveloped to measure the rate of diffusion of molecules across membranes. Thistechnique was applied to DMA and MMA. Permeability coefficients of(5.8 ± 1.0) x 10-8cm s1 and (7.8 ± 2.7) x109 cm s’ at 24°C were measured for the neutral forms ofDMA and MMA, respectively. In addition, the effects ofpH, temperature, and membranecomposition on the diffusion coefficients for DMA and MMA through the liposomalmembrane were investigated. It was determined that only the neutral form of these weakacids permeates across the membrane and that cholesterol decreases the rate ofpermeation of DMA.111TABLE OF CONTENTSAbstract iiList of Tables xList of Figures xiiList of Abbreviations xviiiAcknowledgements xxChapter 1General Introduction 11.1 Speciation and Concentration of Arsenic in the MarineEnvironment 21.1.1 Seawater 21.1.2 Marine Plants and Algae 31.1.3 Marine Animals 61.2 MMA and DMA in the Aquatic Environment 81.2.1 Freshwater 81.2.2 Seawater 81.3 Overview of Thesis 11Chapter 2Exposure of Mussels to[3Hj-MMA 122.1 Introduction 122.1.1 Origins of Organoarsenicals in Marine Animals 122.1.2 Biomethylation of Arsenic 142.1.3 Proposed mechanism for the biosynthesis of arsenosugars 162.1.4 Proposed conversion of arsenosugars to arsenobetaine 18iv2.1.5 Scope of Work 192.2 Experimental 212.2.1 Scintillation Counting 212.2.2 Atomic Absorption Spectrometry 212.2.3 HPLC system 222.24 TLC plates 242.2.5 Chemicals and reagents 252.2.6 The uptake of[3H]-compounds by mussels from seawaterwhich initially contained[3H1-MM.A 252.2.7 Isolation of the[3Hj-labeled arsenicals in the mussel flesh 262.2.8 Identification of the[3HJ-labeled compounds in the musselflesh which eluted from the Dowex column in the NH4Oraction 302.2.9 Analysis of the[311j-labeled compounds in the mussel fleshwhich eluted from the Dowex column in the HC1 fraction 302.2.10 Determination of the total arsenic concentration in the musselflesh, byssal threads and shells 302.3 Results and Discussion 332.3.1 The uptake of[3H]-labeled arsenicals from the seawater 332.3.2 Purification of the[3H]-labeled arsenicals from the musselflesh. 332.3.3 Identification of the[3141-labeled arsenicals in the NH4Ofraction. 362.3.4 Attempted identification of the[3111-labeled arsenicals inthe HC1 fraction. 392.3.5 Total arsenic concentration in the mussel flesh, byssalthreads and shells. 40V2.3.6 Summary 42Chapter 3The biotransformation ofMMA and DMA into arsenobetaine in seawaterand mussels 433.1 Introduction 433.1.1 The origins of arsenobetaine in seawater and marine animals 433.1.2 Scope ofwork 443.2 Experimental 453.2.1 Instrumentation 453.2.2 Chemicals and reagents 453.2.3 The uptake and conversion of[3H1-MMA and[3H1-DMAto[3HJ-arsenobetaine in seawater and mussels. 453.2.4 Isolation and identification of the[3HJ-labeled arsenicals inthe seawater and the mussel flesh. 463.2.5 The diversity of[3Hj-labeled compounds in the seawater 473.2.6 Examination of the[3111-labeled compounds in the seawaterfrom the experiment described in section 3.2.5 473.3 Results and Discussion 483.3.1 Experimental seawater after exposure to[3HJ-MMA and[3H}-DMA 483.3.2 Control seawater after exposure to[3Hj-MMA and[3Hj-DMA 573.3.3 Mussel flesh after exposure to[3H]-MMA and[311]-DMA 623.3.4 The extraction efficiency of the[3Hj-labeled compoundswhich were extracted from the seawater and mussel samples 673.3.5 Examination of the[3Hj-labeled compounds in the seawaterviafter the experiment described in section 3.2.5 683.6 Summary 72Chapter 4The use of liposomes in predicting the biomobility and bioaccumulationofMMAand DMA 734.1 Introduction 734.1.1 How do molecules get across membranes? 734.1.2 Structures of the main phospholipids found in cellmembranes 764.1.3 Structures of hydrated phospholipids 774.1.4 The mobility of the lipid components within the bilayer 794.1.5 Liposomes as models for biological membranes 814.1.6 Types of liposomes 814.1.7 Large Unilamellar Vesicles (LUVs) 824.1.8 How is the biomobility of environmentally sensitivecompounds traditionally modelled? 854.1.9 Difficulties in using partition coefficients to predict thebiomobility of weak acids 854.1.10 The bioaccumulation of arsenicals by Candida humicola 864.1.11 The bioaccumulation ofDMA and MMA by unicellularorganisms 874.1.12 Scope of work and proposed use of diffusion coefficientsacross liposomal membranes as a model for the predictionof biomobiity and bioaccumulation 884.2 Theory 89vii4.2.1 The rate constant, permeability coefficient and activationenergy 894.3 Experimental 914.3.1 Scintillation counting 914.3.2 Chemicals and Reagents 914.3.3 Preparation of LUVs and Sampling Technique 914.3.4 Determination of Octanol/Water partition Coefficients 934.4 Results and Discussion 944.4.1 Rate constants and permeabilities for the diffusion ofMMA and DMA across liposomal membranes (100nm LUVs at pH 7.4) 944.4.2 Activation energy for the diffusion ofMMA and DMAacross liposomal membranes (100 nm LUVs at pH 7.4) 964.4.3 Octanol/Water partition coefficients 984.4.4 Summary 99Chapter 5The Development of an NMR technique to measure the rate of diffusionthrough LUVs and its applications to MMA and DMA 1005.1 Introduction 1005.1.1 Traditional techniques used to measure the diffhsioncoefficients through LUVs 1005.1.2 Spectroscopic techniques used to measure the diffusioncoefficients through LUVs 1015.1.3 Advantages ofusing spectroscopy over traditionaltechniques to measure the diffusion of moleculesacross membranes 1025.1.4 Difficulties in using NIvIR spectroscopy to measure thediffusion of molecules across membranes 1025.1.5 Scope of work 1035.2 Theory 1045.2.1 What causes molecules to diffuse across membranes? 1045.2.2 Diffusion from a spherical vesicle 1065.2.3 The permeability coefficient and activation energy 1095.2.4 Derivation of the equations used to describe the number ofparticles on either side of the membrane 1105.2.5 Permeation of a weak acid 1135.2.6 Comment on D, P, k’, k, ç, kp.j, kMi, and P&j1 (to avoidconfusion later) 1155.3 Experimental 1175.3.1 Instrumentation 1175.3.2 Chemicals and reagents 1195.3.3 Determination of the pK’s for DMA and MMA 1195.3.4 Preparation of LUVs and Sampling technique 1205.3.5 Concepts to consider when designing these experiments 1215.3.6 Bligh-Dyer Extraction 1225.3.7 Phosphorus assay 1235.3.8 Processing the spectra 1235.3.9 Permeability coefficient determination 1345.4 Results and Discussion 1355.4.1 pK determinations for MMA and DMA 1355.4.2 The difii.ision of DMA through LUVs of various sizes 1405.4.3 DMApH study 1415.4.4 MMA pH and temperature study 145ix5.4.5 DMA temperature study 1505.4.6 DMA temperature study with cholesterol in the membrane 1535.4.7 Comments on how the results obtained in this chapterapply to biological samples and possible uses for thetechnique in the future 1575.5 Summary 158Chapter 6The synthesis of organoarsenicals relevant to this thesis 1596.1 Introduction 1596.2 Experimental and Results 1596.2.1 Synthesis of[3H]-MMA from arsenite 1596.2.2 Synthesis of[3Hj-DMA from[3Hj-MMA 1616.2.3 Synthesis of trimethylarsine from arsenic trichloride 1626.2.4 Synthesis of tetramethylarsonium iodide from trimethylarsine 1636.2.5 Synthesis of trimethylarsine oxide from trimethylarsine 1656.2.6 Synthesis of arsenobetaine from trimethylarsine 168Bibliography 170xLIST OF TABLESTABLES PAGE2.1 Graphite Furnace Operating Parameters. 222.2 Operating conditions for the continuous Hydride Generation AtomicAbsorption assembly. 242.3 Rfvalues for the unknown from the “NH4O fraction”. 362.4 The[3H]-containing compound(s) in the “HCI fraction”. 402.5 Arsenic concentrations in the mussel flesh, byssal threads and shells. 402.6 The radioactivity in the shells. 413.1 Percentageof[3H]-labeled compounds which were extracted fromthe seawater and mussel samples. 674.1 Mean diameter of LUVs produced via the extrusion technique. 844.2 Rate constants and permeability coefficients for MMA and DMA atpH 7.4 across the liposomal membrane (100 nm filter size). 944.3 The activation energy for the diffusion of across liposomal membranes. 964.4 The octanol/water parition coefficients for MMA and DMA. 985.1 Raw data from a diffusion experiment. 1255.2 Integral ratios and time for a diffusion experiment. 1265.3 Calculated integral ratios for the diffusion experiment in Table 5.2. 1325.4 pK’s for MMA and DMA 1355.5 The1H-NMR methyl resonance positions for MMA and DMAwith variations in pH. 1385.6 The1H-NMR methyl resonance positions for DMA during a diffusionexperiment. 1385.7 DMA diffusion through different sized LUVs (Temp = 24°C andpH=7.4). 1405.8 DMA pH Study (Temp = 24°C with NaC1 outside). 142xi5.9 DMA pH Study (Temp = 24°C with glucose outside). 1425.10 MMA Diffusion Coefficients (pH = 6.98) through 200 nm LUVs. 1465.11 MMA Diffusion Coefficients (pH = 7.40) through 200 nm LUVs. 1465.12 MMAH Diffusion Coefficients through 200 nm LUVs (from Table 5.10) 1475.13 MMAH Diffusion Coefficients through 200 nm LUVs (from Table 5.11) 1475.14 The activation energy for the permeation ofMMAH through thephospholipid bilayer ofLUVs. 1495.15 Permeability coefficient determination for the anion ofMMA. 1495.16 Rate constant determination for the anion ofMMA. 1505.17 DMA Diffusion Coefficients (pH 7.4) through 200 nm LUVs. 1515.18 DMAH Diffusion Coefficients through 200 nm LUVs. 1515.19 The activation energy for the permeation of DMAH through thephopholipid bilayers of LUVs. 1525.20 DMA Diffusion Coefficients (pH 7.0) through 200 rim LUVs (55:45,EPC:Cholesterol). 1545.21 DMAH Diffusion Coefficients through 200 rim LUVs (55:45,EPC:Cholesterol). 1545.22 The activation energy for the permeation of DMAH through aphospholipid bilayer which contains cholesterol. 156xliLIST OF FIGURESFIGURES PAGE1.1 Formulas and pKs of arsenic containing compounds found in seawater. 21.2 Structures of arsenic containing ribosides found in marine algae. 41.3 Stuctures for arsenic containing phospholipids found in marine algae. 51.4 Some arsenic containing compounds found in marine animals. 61.5 Water column profiles of arsenate (As V), arsenite (As ifi), MMA,DMA, phosphate (HP042),and chlorophyll (CHL) concentrationsduring a bloom in Alice Arm B.C. 92.1 Challenger’s Mechanism for the biomethylation of arsenic. Thearsenic (III) intermediates in braces are unknown. 152.2 Structure of S-adenosylmethionine (SAM). 152.3 Proposed mechanism for the biosynthesis of arsenosugars by marine algae. 172.4 Proposed biosynthetic pathway from arsenosugars to arsenobetaine. 192.5 Schematic diagram of the continuous Hydride Generation AtomicAbsorption assembly. 232.6 A) 200 liter seawater tank with continually circulating seawater;B) 15 liter experimental tank containing[3H]-MMA; C) “controlgroup” of mussels; D) wire cage containing “experimental group”of mussels; E) aeration stones and tubing. 272.7 Schematic representation of the proceedures used for the extraction ofarsenic compounds from the mussels and for the purification of theextract. 292.8 Wet ashing apparatus; (A) Teflon cylindrical plugs; (B) Teflon diffusionfunnels; (C) Teflon stopper with capillary; (D) 250 ml round bottom flask.All joints are 14.5/23. All dimensions are mm. 322.9 The elution profile from a Sephadex LH-20 column for the water soluble[3H]-labeled compouds which were MeOH extracted from the mussels. 342.10 The elution profile from the Dowex column for the[3H]-labeledcompounds which eluted in fraction 8 to 21 from the Sephadex column. 352.11 HPLC chromatograms from the anion exchange column at pH 6.67; (A)arsenobetaine and dimethylarsinate on the anion exchange column atpH 6.67, (B) the compound eluting in fractions 11 to 17 from theDowex column. 383.1 The cation exchange chromatograme for the standards of MMA, DMA,and arsenobetaine. 493.2 The cation exchange chromatograms for the phenol extract from the“experimental seawater” on days 4 (A) and 7 (B) after[3H]-MMAexposure. 503.3 HPLC chromatograms; (A) arsenobetaine, MIvIA, and DMA on the anionexchange column at pH 4, (B) the compound eluting in fraction 3 (Figure3.2 (B)) on the anion exchange column at pH 4. 513.4 HPLC chromatograms; (A) MMA on the size exclusion column and (B)of the compound eluting in fraction 3 (Figure 3.2 (B)) on the size exclusioncolumn. 523.5 HPLC chromatograms; (A) arsenobetaine and DMA on the anion exchangecolumn at pH 6.67, (B) the compound eluting in fraction 7 (Figure3.2 (B)) on the anion exchange column at pH 6.67. 533.6 HPLC chromatograms; (A) arsenobetaine on the size exclusion column,and (B) of the compound eluting in fraction 7 (Figure 3.2 (B)) on the sizeexclusion column. 543.7 The cation exchange chromatograms for the experimental seawater on days4 (A) and 7 (B) after[3HJ-DMA exposure. 563.8 The HPLC chromatogram for the compound eluting in fraction 7 (Figurexiv3.7 (A)) on the anion exchange column at pH 6.67. 573.9 The cation exchange chromatograms for the phenol extract from the“Control seawater” on days 4 (A) and 7 (B) after[3H]-MMA exposure. 583.10 The HPLC chromatogram for the compound eluting in fraction 7 (Figure3.9 (B)) on the anion exchange column at pH 6.67. 593.11 The cation exchange chromatograms for the phenol extract from the“Control seawater” on days 4 (A) and 7 (B) after[3H]-DMA exposure. 613.12 The I{PLC chromatogram for the compound eluting in fraction 7 (Figure3.11(B)) on the anion exchange column at pH 6.67. 623.13 The cation exchange chromatograms for the phenol extracted compoundsfrom the mussel flesh on days 4 (A) and 7 (B) after[3Hj-MMA exposure. 633.14 The HPLC chromatograms for the compound eluting in fractions 2 and 3(Figure 3.13 (B)) on the anion exchange column at pH 4. 643.15 The cation exchange chromatograms for the phenol extracted compoundsfrom the mussel flesh on days 4 (A) and 7 (B) after[3H]-DMA exposure. 663.16 The size exclusion HPLC chromatogram for the “experimental seawater”on days 4 (A), 7 (B), and 10 (C). 713.17 The size exclusion HPLC chromatograms for the DMA standard. 714.1 The Fluid Mosaic model of the cell membrane. The arrows represent thedirections of the lateral diffusion of the proteins. 744.2 Mechanisms of permeation through the bilayer without entry into thehydrophobic region. 754.3 Structures of the major phospholipids found in cell membranes; PA,phosphatidic acid; PS, phosphatidyl serine; PC, phosphatidyl choline;PE phosphatidylethanolamine; P1, phosphatidyliniositol; DPG,diphosphatidyiglycerol. R is the side chain (Figure 4.4) 76xv4.4 The structures of the side chains of egg phosphatidyl choline andpalmitic acid (a), stearic acid (b), oleic acid (c), and linoleic acid (d). 774.5 Structures of phospholipids which are formed upon hydration; A) amonolayer formed at an air-liquid interface; B) a micelle; C) an example ofa liposome; D) a black membrane, a planar bilayer, that separates twoaqueous phases. 784.6 Modes of mobility of the lipid components of the bilayer. 794.7 Insertion of cholesterol into a phospholipid bilayer. A schematicrepresentation of the alignment of a cholesterol molecule with aphospholipid in a lipid bilayer. 804.8 The Extruder. 824.9 Electron micrographs of vesicles prepared by extrusion through; A) 400;B) 200; C) 100; D) 50; E) 30 nm pore size polycarbonate filters. 834.10 Test of the unilamellarity of vesicles extruded the indicated number oftimes through 400 (.), 200 (a), and 100 () nm pore size filters. 844.11 Radiolabeled sampling proceedure. 924.12 Efflux measurements of MMA and DMA. Data is plotted accordingto equation 4.6. 954.13 Activation energy determinations for MMA and DMA. Data is plottedaccording to equation 4.8. 975.1 Flux across a membrane in response to a difference in chemical potential. 1055.2 Schematic depiction Ofvaq1,vaq0, vmout, D, K, k’, k,[pAjm0 [HAjm, [JTA]aq0 [HAJaq. 1075.3 Schematic depiction OfCm1,C, rj, and rout. 1085.4 The two chemical species of DMA in aqueous solution. 1135.5 The pulse sequence used for water suppression. 1175.6 The microprogram designed to operate the spectrometer. 118xvi5.7 The ‘H-NMR spectrum of DMA and LUVs. 1245.8 An example of a plot of the inside (ii) and outside (+) integral ratiosversus time for the diffusion ofDMA through a liposomal membrane. 1275.9 An example of a plot oflfl(nt1neCl) (ii) and (.÷)versus time for the diffusion of DMA through a liposomal membrane. 1305.10 An example of the inside (‘I) and outside (÷) integral ratios versus timefor the diffusion of DMA through liposomal membranes. The solid linesare the experimental fits for the data which were generated by usingequations [5.471 and [5.48]. 1335.11 A plot of the LHN14R methyl resonance positions for MMA versus pH. 1365.12 A plot of the1H-NMR methyl resonance positions for DMA versus pH. 1375.13 A plot of the lH..NTvII?,. methyl resonance positions for DMA versus timefor a diffusion experiment. 1395.14 A plot of the pH dependant rate constants (k in ml s) vs the [H+] forthe efflux of DMA at 24°C. Either glucose (a) or NaCI (.)was added tothe outside of the 200 nm LUVs to balance the osmotic pressure. 1435.15 A plot of the pH dependant rate constants (Ic in s1) vs the [J.J+] forthe efflux of DMA at 24°C. Either glucose (a.) or NaCl () was addedto the outside of the 200 nm LUVs to balance the osmotic pressure. 1445.16 A plot of the ln(P1sjj) from the MI{AH dififusion experiments at pHs6.98 () and 7.4 () versus lIT. 1485.17 A plot of the ln(Ppjj) versus l/T for the activation energy determinationfor DMAH diffusion through a phospholipid bilayer. 1525.18 A plot of the ln(PM.1) versus l/T for the activation energy determinationfor DMAH diffusion through a phospholipid bilayer which containscholesterol. 1556.1 ‘H-NMR (200MHz in D20) spectrumof[3Hj-MMA. 160x’16.2 ‘H-NMR (200MHz inD20) spectrumof[3H1-DMA. 1626.3 ‘H-NMR (200MHz in D20) spectrum of tetramethylarsonium iodide. 1646.4 DCI mass spectrum of tetramethylarsonium iodide. 1656.5 1H-NMR (200MHz in D20) spectrum of trimethylarsine oxide. 1666.6 DCI mass spectrum of trimethylarsine oxide. 1676.7 1H-NMR (200MHz in 020) spectrum of arsenobetaine. 1686.8 DCI mass spectrum of arsenobetaine. 169xviiiLIST OF ABBREVIATIONSACS American Chemical Societycm centimeterDCI-MS desorption chemical ionization-mass spectrometryDMA dimethylarsinate and/or dimethylarsinic acidDMA dimethylarsinateDMAH dimethylarsinic acidg gramHPLC high performance liquid chromatographykg kilogramL literLUVs large unilamellar vesiclesM molar (mol L1)mA milliamperemCi millicuriemg milligramml millilitermm millimetermM millimolarMMA monomethylarsonate and/or monomethylarsonic acidMMA monomethylarsonateMIvIAH monomethylarsonic acidnm nanometerppb parts per billionppm parts per millionpsi pounds per square inchrpm revolutions per minutes secondTSP 3-(Trimethylsilyl) proprionic-2,2,3,3-d4sodium salt• jCi microcuriemicrogramvs versusv/v volume-to-volume ratiow/v weight-to-volume ratioxixxxACKNOWLEDGEMENTSI would like to extend my sincere gratitude to my research supervisors, Dr. W.R.Cullen and Dr. F.G. Herring, for their guidance, patience, encouragement, and financialsupport throughout the course of this project.I would also like to thank Dr. P.R. Cullis and K. Wong for allowing me to performthe experiments described in Chapter 4 in their lab.In addition, Iwould also like to thank Dr. P.S. Philips for helping me with themathematics behind the diffusion equations and Dr. 0. Chan for helping me with the NIvIRspectrometer.I would also like to acknowledge the members of my research group, both pastand present, for many helpful discussions.CHAPTER 1GENERAL INTRODUCTIONArsenic is the 20th most abundant element in the earth’s crust.’ Its averageconcentration is often quoted as 3 ppm (parts per million), but this may vary widelydepending upon geographical location.2,3Arsenic is generally associated with igneousand sedimentary rocks4 and in particular, with the sulfidic ores of realgar (As4S),orpiment (As2S3)and arsenopyrite (FeAsS). Arsenic is released into the environment bynatural processes such as weathering, volcanic and biological activity, and byanthropogenic activities such as the smelting of ore and the burning of fossil fuels.5’6Arsenic is present in virtually all types of soils, sediments, aquatic environmentsand organisms.5 In particular, it is present in high concentrations (up to 100 mg/kg ) inmarine algae and animals.7 Since marine organisms are a major source of arsenic inhuman beings, 7 information about its chemical forms is important from toxicological andnutritional points of view.The literature on arsenic in the sea dates back to the beginning of the century. 8-10By the 1920’s it was firmly established that marine organisms contained high levels ofarsenic.”12 In spite of much effort, however, the chemical form of arsenic in marineorganisms remained unknown until 1977, when arsenobetaine 1 was isolated and identifiedin the tail of the Western rock lobster.’3 Arsenobetaine has since been identified invarious marine animals and the ubiquitous nature of this arsenical in marine animals hasbeen well established.5 In addition to arsenobetaine, numerous other arsenic containingcompounds have been isolated and identified in marine organisms.Each of the individual arsenic containing compounds has different physical andchemical properties, toxicities, mobilities and biotransformations. In order to fullyunderstand the arsenic biogeochemical cycle in marine organisms, the differentcharacteristics of each of these compounds must be throughly investigated.521.1 SPECIATION AND CONCENTRATION OF ARSENIC CONTAININGCOMPOUNDS IN TILE MARINE ENVIRONMENT1.1.1 SeawaterIn seawater the total arsenic concentration is typically 1 to 8 ppb (parts perbillion).147 Arsenate 2 and arsenite 3 are the predominant arsenic species in seawaterand their concentrations depend upon the nature and degree ofbiological activity at thetime of sampling and the redox potential of the seawater.’8 The ratio of arsenate toarsenite in seawater is typically about 10:1. Both phytoplankton and bacteria can reducearsenate to arsenite.’9 In addition to the inorganic arsenic species, two methylatedarsenicals, monomethylarsonate (MMA) 4 and dimethylarsinate (DMA) 5 are also presentas a small percentage of the total arsenic concentration in seawater.20-2 Arsenobetaineis identified in seawater as part of this thesis. Arsenate, MMA, DMA and arsenobetaineall contain arsenic in an oxidation state of(V), whereas in arsenite and trimethylarsine theoxidation state is (III).+(CH3)AsCH2COO H3AsO4(1) pK4.6 (2) pK1=2.2pK=6.9pK311.5As(OH)3(3) pK1 = 9.2 (CH3)AsO(OH)2(4) pK14.11pK28.77(CH3)2AsO(OH)(5) pK6.28Figure 1.1 Formulas and pKs of arsenic containing compounds found in seawater.31.1.2 Marine Plants and AlgaeArsenate is typically found in marine plants and algae,23 however, the details ofthe uptake mechanism are poorly understood. Arsenate is either accumulated by algae due..to its similarity with phosphate22’4 5 or absorbed by a non-competitive mechanism.26Regardless of the mechanism of arsenate uptake, however, the primary producers such asthe phytoplankton and algae must adapt for the accumulation of cellular arsenate. Themarine phytoplankton (diatoms, coccolithophorids, dinoflagellates, and green algae) haveadapted to cellular arsenate by converting it to arsenite, MMA and DMA which are thenexcreted from the cells.22 The metabolism of arsenic species depends upon the type ofmarine phytoplankton, but in general 30 to 50% ofthe arsenic which is not excreted isconverted to lipid soluble arsenic compounds.The three main classes of macroalgae contain arsenic concentrations between 0.4 -32 ppm.27 The classes and percentages of arsenic in the organic form are as follows:Phaeophyceae, 78%; Rbodophyceae, 57%, and Chlorophyceae, 53%. Theorganoarsenicals in macroalgae were first isolated and identified by Francesconi andEdmonds.28’9 They extracted these organoarsenicals from the seaweed Eckloniaradiara, and used chromatographic and spectroscopic techniques to show that the algalarsenic was incorporated into both water soluble and lipid soluble compounds. The watersoluble compounds were found to be the arsenic containing ribosides 6a and 6b andaccounted for 81% ofthe total arsenic which was present in the seaweed. The arseniccontaining ribosides 6c and 6d were subsequently isolated from the kidney of the giantclam Tridacna maxima,3°these organoarsenicals were attributed to the symbiotic,unicellular, green algae which lives in the clam tissue.30’ These arsenic containingribosides along with 6e have since been shown to be present in many other marineseaweeds.2830’32 6 Recent studies on the macroalgae Sargassum thurbegii haveshown the presence of 6f the trimetbylarsonium analogue of 6e, as a minor arsenical.3740(CH3)2As(6)a R =CH2HO CHOb R =CHHO CHS3c R =CHOHCHOPO(O )OCHd R =CH2HO CHOSO3e R=CHCHM1HSO(CH3)As(61)ORHO OHFigure 1.2 Structures of arsenic containing ribosides found in marine algae.5Two of the structures34for the arsenic containing phospholipids are 7a and 7b.The phosphorus containing arsenical 7c has been identified in E. rathata28 and Hizikiafusforme.39 It has been speculated that arsenic containing phospholipids may bebiosynthesized by the algae to facilitate the incorporation of arsenic into the membranelipid bilayers which will ultimately result in its excretion from the algae.31The ability of algae to tolerate high levels of arsenic is exemplified by the the greenflagellate, Tetraselmis chuii, which survives in arsenic concentrations as high as 1000 ppmarsenate.4°(The average concentration of arsenic containing compounds in marine algaeis about 10 ppm.5)0 0‘I(CH3)2As OCH2CHOHCHPOROH(7)a R =CH2H(OOCX)CHO Xb R =CHH(O )CH00CXc R =CH2H(O )CHOFigure 1.3 Structures for arsenic containing phospholipids found in marine algae.61.1.3 Marine AnimalsThe arsenic concentration in marine animals varies widely between 0.31 ppm foundin the flesh of a typical salmon to about 340 ppm found in the midgut of the carnivorousgastropod Charonia sauliae.5 The arsenic containing compounds in marine animals arerelatively non-toxic. Crustaceans and other benthic organisms have higher arsenicconcentrations than fish. Since its identification from the tail of the Western rocklobster,13 arsenobetaine has been shown to be the predominant arsenical in most marineanimals.5’283241 When arsenobetaine was injected into mice at concentrations of up to500 mg kg, it was rapidly excreted without being altered metabolically.42 The mice inthese experiments showed no symptoms of poisoning. Arsenobetaine has also beenisolated from the urine of human subjects who had eaten cooked lobster tails andplace.43’4 Additional feeding experiments with arsenobetaine and humans showed that69-85% of the ingested arsenic was excreted within 5 days.44In addition to arsenobetaine, other methylated arsenicals have been identified inmarine animals. The tetramethyl arsonium ion 8 has been found in the clam Meretrixlusoria45 and in several gastropods.46’7 There have been claims of the presence ofarsenocholine 9 in shrimp,485°however, its presence could not be confirmed byothers.50’1 Arsenocholine 9 is generally believed to be a probable precursor toarsenobetaine.52 Trimethylarsine oxide 10 has been found as a minor component of fishand some molluscs.47’5355Shibata and Morita have recently shown the presence of the arsenic containingribosides 6a and 6c in a wide variety ofbivalves,56by using high performance liquidchromatography coupled to an inductively coupled argon plasma atomic emissionspectrometer as the detector. They speculate that these arsenic containing ribosides enterthe bivalves via their food, i.e., the phytoplanktons are the origin of these sugars. The• quantity of the arsenic containing ribosides varies widely even within the same species.7+(CH3)4As (CH3)AsCH2CO(8) (9)(CH3)AsO(10)Figure 1.4 Some arsenic containing compounds found in marine animals.An interesting feature of the arsenic compounds in marine animals is that, with theexception of the arsenic containing ribosides found in marine animals, all of the nitrogenanalogues of the arsenic compounds are abundant in these organisms. For example,glycinebetaine, tetramethylammonium ion, and trimethylamine oxide are the nitrogenanalogues of arsenobetaine, tetramethylarsonium ion, and trimethylarsine oxide,respectively.7 These nitrogen containing compounds are thought to be the end productsof the nitrogen metabolism, and are probably retained in the body as osmo-regulators.57The nitrogen containing compounds are present in much greater quantities than theirarsenic analogues.781.2 MMA AND DMA IN THE AQUATIC ENVIRONMENT1.2.1 FreshwaterThe arsenic concentration in fresh water environments is typically 1-10 ppb,2although values as high as 3500 ppb have been reported.58 Braman and Foreback59firstdetected MMA and DMA in aquatic systems in 1973. They reported on the arsenicspeciation in both freshwater and saline samples. The freshwater samples containedarsenicals, arsenate and usually arsenite. MMA and DMA were also usually present,typically between 10 and 15% of the total arsenic, but values up to 70% were reported. Inall cases the dominant organoarsenic compound was DMA. A similiar study by Andreaeon the distribution of the organoarsenicals in fresh water suggested that there is acorrelation between the sites with the highest density of planktonic algae and the highestconcentration of methylarsenicals.6°Arsenite is methylated by various freshwater species of green algae, includingAnkistrodesmus sp., Chiorella sp., and Selenastrum sp. Each of these organismstransformed arsenite at a concentration of 5000 ppb to MMA, DMA and trimethylarsineoxide.6’rvtA6266 and DMA67 are used as herbicides and, therefore, it may be expectedthat MMA and DMA would find their way into the surrounding lakes and rivers.Fortunately, both MMA and DMA are highly retained by the soils and hence leaching intofresh water does not readily occur.651.2.2 SeawaterAndreae has also reported detailed concentration versus depth profiles for MMAand DMA in northeast Pacific and Californian coastal waters.60’8 The general trend inthese profiles seems to be a depletion of arsenate concentration in the photosyntheticallyactive surface waters (eutropic zone) with a corresponding increase in arsenite, MMA, and9DMA concentrations. The DMA to MMA concentration ratios in these profiles wastypically about 10:1. The concentration ofDMA decreased towards the base of the photiczone while the concentration ofMMA remained fairly constant. In deeper water themethylarsenicals were either below the detection limits or barely detectable. Thisobservation suggests that DMA and MMA were produced by phytoplankton or closelyrelated heterotrophs. This idea is fhrther supported by correlations which were foundbetween typical indicators of primary productivity, i.e., chlorophyll concentrations6°and/or 14C uptake.68 It was concluded from these studies that arsenate was taken up byphytop!ankton in the eutrophic surface waters and subsequently converted to arsenite,MMA and DMA which were then released back into the water column. The above trendsare prevelant in the concentration versus depth profiles in Figure 1.5. The sharp decreasein DMA concentration with depths below 20 m (Figure 1.5) would suggest release ofDMA from live algal cells rather than bacterial decomposition of organoarsenical, as thelater process would be expected to distribute these compounds to a greater depth.Other studies have shown how the type of algal species17 and temperature697°affects the appearance, concentration and relative proportions of these twomethylarsenicals.10Ia12$.175As (V) (ppb)As (III), MMA, and DMA (ppb).ie .zeAs(Ill)DMAFigure 1.5 Water column profiles of arsenate (As V), arsenite (As ifi), MMA, DMA,phosphate (HPO42),and chlorophyll (CHL) concentrations during abloom in Alice Arm B.C.5e2SSe75CHLHP042eeHP042(p.g L1) and CHL (mg m3)20:‘MMAaJaIIIIIIa6080iee111.3 OVERVIEW OF THESISThe work described in this thesis was initiated so that the role ofMMA and DMAin the marine biogeochemical cycle of arsenic could be better understood. These twomethylarsenicals are the simplest organoarsenicals found in the aquatic environment and,therefore, MMA and DMA may be likely precursors to many of the more complexorganoarsenicals. Thus, the results from this work should provide information about themetabolic pathways of arsenic in the marine environment. In addition, different amountsof these two very similiar compounds are bioaccumulated by marine organisms and arefound in seawater. This is quite possibly the result of differences in the ability ofMMAand DMA to pass through biological membranes.The biotransformations of MMA and DMA were investigated by adding either[3Hj-MIvIA or[3H]-DMA to a seawater tank. This seawater tank also contained musselsand at the end of the exposure period the[3}1]-labeled arsenic compounds in the musselsand seawater were identified or an attempt was made to identi1’ them. In addition, thetotal arsenic concentrations in the mussel flesh, byssal threads and shells were determined.The diffusion coefficients of MMA and DMA across phospholipid membraneswere determined by using a radiolabeled permeant and by using a NMR technique whichwas developed as part of this thesis. These coefficients are suggested as a way to predictbiomobility and the disadvantages ofusing partition coefficients for such predictions isdiscussed. The effects of temperature, membrane composition, and pH on the diffusioncoefficients ofMMA and DMA across membranes were investigated.The synthesis of many of the organoarsenicals which were used in this thesis is alsodescribed, including the synthesis of radiolabeled[3H1-MMA and[3Hj-DMA.12CHAPTER 2EXPOSURE OF MUSSELS TO[3H]-MMA• 2.1 INTRODUCTION2.1.1 Origins of the Organoarsenicals in Marine AnimalsIt is generally assumed that the organoarsenicals which are found in marineorganisms are partially synthesized from arsenate by organisms at lower trophic levels.5This hypothesis is based upon the observation that species at the higher trophic levels(fish, crustaceans, and gastropods) do not seem to be able to synthesize compounds suchas arsenobetaine from arsenate.The accumulation hypothesis is supported by Unlu and co-workers71 73 whoshowed that[74As]-algal organoarsenicals (presumably compounds 6a-f) were readilytaken up by crabs, whereas inorganic[74As]-arsenate, fed via arsenate injected musselswas not readily taken up, and the74As]-arsenate that was taken up was readilyeliminated from the crab without being converted to any[74As]-organoarsenicals.Francesconi and Edmonds injected earthworms with arsenate which were hand-fed tocatfish and school whiting. Very little of the ingested arsenate was retained by the fish andthe amount of arsenobetaine in the fish did not increase over the controls.53 However,trimethylarsine oxide was detected in the dosed whiting and the catfish showed an increasein this arsenical relative to the control. It was argued that arsenate was not converted totrimethylarsine by the fish, but that the biotransformation was the result of bacterial actionin the gut tract of the fish.53The direct transfer of organoarsenicals along the food chain is not a universalphenomenom. The organoarsenicals found in algae are clearly not the same compounds13found in the animals which feed on them as shown by the following example. The algaeDunaliella tertiolecta was cultured in the presence of[74As]-arsenate and fed to thelobster Homarus americanus. The results of a chromatographic study showed that no[74As]-arsenobetaine was produced during the time scale of the experiment (2 days). Inaddition, the arsenic containing phospholipids 7a,b,c were degraded to the simple arseniccontaining ribosides, such as 6a.74 Thus, the lobster appears to be unable to convert thearsenic containing ribosides into arsenobetaine, its natural organoarsenical. When[74AsJ-arsenate is fed directly to the lobster, no organoarsenicals seem to be produced.Subsequent experiments yielding similiar results have been tried with gastropods.75However, Klumpp and Peterson75-7have obtained experimental results whichseem to contradict the generality of the accumulation hypothesis. In the marine foodchain, Fucus spiralis—Littorina littoralis—Wucella lapillus, algae are being consumed bya herbivorous snail which is in turn being consumed by a carnovorious snail. L. littoralisincorporates[74As]-arsenate from the water or the[74As]-labeled F. spiralis, whereas, Nlapillus incorporates much less[74As]-arsenate from water. Both snails are equallyefficient of accumulating arsenic from their food and this is their main source of theelement. The majority of the[74As] in the snails, regardless of the feeding mechanism,seems to be a single water-soluble compound which is not present at the first trophic level.Some of the lipid-soluble arsenical found in F. spiralis is present in both snails at about10% of the label. These results indicate that the principal water soluble arsenical in thesnails does not come from the algae and that the snails can synthesize it directly fromarsenate. Furthermore, the lipid soluble arsenic compound is passed along the food chain.In general, the arsenic concentration in herbivorous species is lower than in carnivores.7880Also, the arsenic concentration in a marine animal is not necessarily the same asthe arsenic concentration in the surrounding environment. For example, the arsenic14content in clams is certainly not dependent upon the arsenic content of the sediments inwhich they are living. This was shown by the chalky Macoma (Malcoma calcarea) whichwhen living in the mine-polluted sediment (46.5 ppm As wet weight) was observed tocontains 3.2 ppm As and when living in cleaner sediment (9.4 ppm As) contains 3.9 ppmAs.52.1.2 Biomethylation of ArsenicThe products of the biomethylation of inorganic arsenic have been observed inmicroorganisms, plants, mice, monkeys and humans.8’ It has been suggested thatorganisms biomethylate arsenic as a method to detoxif,’ it.82The biomethylation of arsenic was first described in detail by Challenger after hecorrectly identified the volatile compound produced by the mould Scopulariopsisbrevicaulis as trimethylarsine.83 Trimethylarsine had a garlic like odor and had beenlinked to several cases of arsenic poisoning.82’4 Challenger proposed a mechanism forthe biotransformation of arsenate to trimethylarsine82(Figure 2.1). This scheme involvesalternating reduction and oxidative steps with the transfer of a methyl group to the arsenicoccuring during the oxidation step. Support for this sequence comes from the observationthat arsenite, arsenate, Mv1A, and DMA are all substrates for the production oftrimethylarsine by S. brevicaulis.Challenger suggested that the methyl donor was probably a methylated compoundsuch as betaine, methionine or a choline derivative. Of these three suggestions he foundthat only[14CH3]-methionine transferred its label to arsenite to any significantextent.85’6 The active form of methionine has subsequently been identified as Sadenosylmethionine (SAM)87’8(Figure 2.2). This has been supported by Cullen eta!. 89-91 who showed that the CD3 group from labeled L-methionine-methyl-d3152e (CH3) 2eH3AsO4 -* As(OH)3 -* CH3AsO(OH)2 -* {CH3As(O )}(2) (3) (4)2e (CH3) 2e(CH3)As (CH3)AsO— {(CH2As(OH)} -(CH3)AsO(OH)(11) (10) (5)Figure 2.1 Challenger’s Mechanism for the biomethylation of arsenic. The arsenic(III) intermediates in braces are unknown.0430% I—GI— 2 2 S—HO INHOH OHFigure 2.2 Structure of S-adenosylmethionine (SAM)16was tranferred intact to a wide variety of arsenicals by Scopulariopsis and Candidacultures.Investigations into the identity of the reducing agent in the scheme have beencarried out92 and a range of thiols and dithiols including: cysteine, glutathione,dithiothreitol, and lipoic acid were found to be capable of canying out the reductions via atwo electron transfer process.9293The first well documented report of an arsine being produced by bacteria was notmade until 1971, when an anaerobic strain ofMethanobacterium produced a volatile gaswith a garlic odor from arsenate.94 This volatile gas was probably dimethylarsine.Carbon dioxide was necessary for its production. Both dimethylarsine and trimethylarsinewere produced by cell extracts ofMethanobacterium thermoautotrophicum.95Since 1971, a number of nonmethanogenic bacteria have been identified byAnderson and co-workers as methylarsine producers.96-8 The bacteria that were used intheir studies were isolated from the environment and cultured in media containing arsenate(100 ppm). Under aerobic conditions arsenate and MMA were methylated by thebacteria to dimethyl and trimethyl species. In addition, the bacteria also reduced MMA tomethylarsine and demethylated MMA to arsenate.The occurence ofMMA, DMA, and trimethylarsine oxide in the sediments,seawater, and selected marine organisms and plants seems to support Challenger’smechanism (Figure 2.1) of biomethylation.2.1.3 Proposed mechanism for the biosynthesis of arsenosugarsFrancesconi and Edmonds have proposed the following scheme, Figure 2.3, toaccount for the biosynthesis of arsenosugars by marine algae.99J0°The scheme beginswith the incorporation of arsenate by the algae, which may be accumulated because of itssimilarity to phosphate. The arsenate is then presumably metabolized by the170IIOHMethylation0Methylation H> Me2As-O-Methylation7?0HMe3AsFigure 2.3 Proposed mechanism for the biosynthesis of arsenosugars by marine0IIMeAs-OOHAdenosylation ‘ Adenosylation..,NH2HO(12)GlycosidationMethylation•0—7?MethylationGlycosidationtORalgae.9918mechanisms outlined by Challenger (Figure 2.1), to initially produce MMA and thenDMA. It would also seem plausible to start the scheme with either DMA or MMA, bothofwhich are present in seawater in varying amounts.5 The arsenic containing nucleoside12 is produced if S-adenosylmethionine (Figure 2.2) donates an adenosyl group to DMAinstead of a CH3 group. The nucleoside 12 has recently been isolated by Edmonds eta!.from the kidney of the giant clam, Tridacna maxima.’°’ This nucleoside (uponenzymatic hydrolysis) could yield the range of dimethylarsinoribosides which have beenidentified from algae sources. A possible biosynthetic pathway to the formation of thetrimethylarsinoriboside is also shown in Figure 2.3 and the incorporation of a third methylgroup attached to the arsenic may occur at three possible steps.2.1.4 Proposed conversion of arsenosugars to arsenobetaineFrancesconi and Edmonds have suggested that the arsenic containing ribosideswhich are found in algae are the likely precursors ofarsenobetaine.99’10°The proposedroutes are shown in Figure 2.4. Anaerobic decomposition experiments on the naturallyoccuring arsenic containing ribosides which are found in the kelp E. radiata provideevidence to support this hypothesis. Anaerobic conditions occur in marine sediments andthe reactions may be microbially mediated. Clevage of the C3-C4 bond in the ribose ringof the dimethylarsenoribosides yields dimethyloxarsylethanol, and cleavage of the samebond in the trimethylarsenoriboside yields arsenocholine. Both dimethylarsenoylethanol13 and arsenocholine were produced from the appropriate precursors in the experiments.The trimethylarsenoriboside is only a minor constituent (5%) of the totalorganoarsenicals in algae, therefore, both possible pathways to the formation of19arsenobetaine will be considered. Dimethyloxarsylethanol would require a methylation ofthe arsenic and the oxidation of the primary alcohol through intermediates such asdimethylarsinylacetic acid 14 or arsenocholine. Arsenocholine is the more likelyimmediate precursor to arsenobetaine as its presence has been reported inshrimps49’50102and it is readily converted to arsenobetaine when administered to thejuvenile yelloweye mullet. 103anaerobic decomposition oxidation6a-e -* -> -* — -*(CH3)2AsCHO --*(CH3)2AsCO.1- (13) ..L. (14)1 methylation 4’ methylation6f —> —> —> -—*(CH3)AsC2CHO —> —>(CH3)AsC2COanaerobic decomposition oxidation(9) (1)Figure 2.4 Proposed biosynthetic pathway from arsenosugars to arsenobetaine.2.1.5 Scope of workThe objective of the present study is to investigate the uptake of[3}{]-labeledarsenicals by mussels after they had been exposed to seawater which initially contained[3Hj-MMA. After exposure, the[3H]-labeled arsenicals in the mussel flesh wereidentified or an attempt was made to identify them. The total arsenic concentration in the20mussel flesh, byssal threads and shells were determined before and after exposure to[3H]-MMA.Mussels were chosen for this study because they are readily available, easilymaintained and globally distributed. They are known to contain arsenobetaine,arsenosugars, and tetramethylarsonium ion. Mussels are filter feeders and phytoplanktonand diatoms form the bulk of their diet. 104212.2 EXPERIMENTAL2.2.1 Scintillation countingA Packard Tricarb 1900 TR liquid scintillation counter was used to measure the[3H]-activity of chromatographic fractions, extracts, and wipe tests. Packard softwarewas used to run the counter which was standardized daily by using Packard standard #22which contained 0.2 iiCi of[3H]. The scintillation counter was coupled to an EpsonLX-8 10 printer.Typically, between 100 to 300 p.1 of sample or a strip of TLC sheeting (1cm x 3cm) was added to a 7 ml glass scintillation vial which contained 5 ml of scintillationcocktail. The samples were counted for three minutes each and then recounted. Theresults for each sample were averaged and tabulated.2.2.2 Atomic Absorption SpectroscopyA Varian Techtron Model AA 1275 single beam atomic absorption spectrometerwas used to determine the arsenic concentrations in the samples. The spectrometer wasfitted with a Varian Spectra AA hollow cathode lamp which operated at 7 mA. Theatomic absorption signal was monitored at a wavelength of 193.7 nm for all arsenicdeterminations. The spectrometer was equipped with a deuterium background correctorand a GTA-95 accessory for graphite furnace atomic absorption (GFAA) spectroscopy.GFAA spectroscopy was used as the detection method for standards which were run onthe various types of chromatographic systems employed in this work. The sample wasinjected into a pyrolytically coated graphite tube in the furnace by using the automaticdelivery system of the accessory. It was mixed with 20 p.1 of palladium nitrate (100 ppmin 2% w/v citric acid) modifier prior to injection. The optimized furnace operating22Table 2.1 Graphite Furnace Operating Parameters.Step No. Temp. (°C) Time (s) Gas flow Read Comment(L mlir1) Command1 85 5 3.0 No dry2 95 40 3.0 No dry3 120 10 3.0 No dry4 1400 6 3.0 No ash5 1400 2 0 No ash6 2400 0.7 0 Yes atomize7 2400 2 0 Yes atomize8 2400 2 3.0 No cleanparameters are given in Table 2.1. The graphite furnace could be removed and replacedby a hydride generation apparatus for hydride generation atomic absorption (HGAA)spectroscopy. HGAA spectroscopy was used for the determination of the total arsenicconcentrations in the seawater and in the acid digested mussel flesh, byssal threads, andshells. The continuous flow hydride generation assembly is illustrated in Figure 2.5. Itconsisted of a Gilson Miniplus 2 four channel peristaltic pump which was used towithdraw the sample and mix it with hydrochloric acid or buffer solution. This solutionwas then reacted with sodium borohydride solution (2% w/v) and introduced into the gas-23liquid separator via a 20 turn mixing coil. The gases were led into an open-ended Tshaped quartz curvette (8.5 cm x 1 cm (o.d.)) mounted in the air/acetylene flame of astandard Varian burner. The operating conditions which were previously established byCullen and Dodd46 are shown in Table 2.2.QuartzAbsorptionCellSampleAcidNaBH4NitrogenFigure 2.5 Schematic diagram ofthe continuous Hydride Generation AtomicAbsorption assembly.ReactionCoilPeristalticPumpTransferTubeGas / LiquidSeparatorNeedleDrain ValvesPressureRegulator24Table 2.2 Operating conditions for the continuous Hydride Generation AtomicAbsorption assembly.Uptake flow Sample 7.5 ml niiiriHC1 2.0 ml min1NaBH4 4.0 ml min1Carrier gas flow Nitrogen 0.6 L mindHCI concentration 4 MNaBH4concentration 2% (w/v) in 0.1% (w/v)NaOH solution2.2.3 HPLC systemThe HPLC system consisted of a Waters M45 pump, a Waters U6K injector, aWaters column (either a Protein Pak DEAE column 7.5 mm (ID) x 7.5 cm or a ProteinPak 300SW column 7.5 mm (ID) x 30 cm) and a Gilson Micro Fractionator automatedcollector. The eluants were degassed and filtered through 0.5 .im pore size filters fromMillipore. Prior to injection the samples were filtered through 0.5 jim pore size cartriagefilters from Millipore. The collected fractions were manually transferred to either thescintillation counter or the atomic absorption spectrometer for detection.2.2.4 TLC platesThe TLC plates were aluminum backed, precoated with 0.2mm of silica gel, andhad dimensions of 20 x 20cm.252.2.5 Chemicals and reagentsAll chemicals used from BDH and Fisher Scientific were classified as analyticalgrade and meet ACS specifications, respectively. The solvents which were used wereobtained from Fisher Scientific and were classified as HPLC grade. A standard arsenicsolution (1000 ig m11 ) was prepared by dissolving 1.3203 g of arsenic trioxide (As203,Fisher Scientific) and 2 g ofNaOH in 20 ml of deionized water. The solution was dilutedto 200 ml, neutralized with HC1 (12 M) and made up to 1000 ml. This solution wasdiluted as necessary to prepare working solutions for GFAA and HGAA spectroscopy.Deionized water (Aquanetics Aqua Media System) was used for solutionpreparation and in all of the liquid chromatography. All glassware and plasticware thatwere not used with the[3H]-labeled compounds were cleaned by soaking overnight in 2%(v/v) Extran solution, rinsing with water and soaking in HC1 (1 M) overnight. All of theglassware and plasticware that were used with the[3H]-labeled compounds were rinsedwith water, soaked overnight in a 5% (v/v) solution of Count-Off. After the overnightsoakings the utensils were first thoroughly rinsed using tap-water and then deioinizedwater.The synthesis of the[3HJ-MMA,[3H]-DMA, arsenobetaine, trimethylarsine oxideand tetramethylarsonium ion used in this chapter is described in Chapter 6.2.2.6 The uptake of[3HJ-compounds by mussels from seawater which initiallycontained13H1-MMAApproximately 150 mussels (Mytilus edulis), ranging in shell length from 2.5 to5.0 cm, were collected from the beach surrounding Point Grey in Vancouver B.C. Thesewere divided into two groups of about 75 mussels each. The first group was designated26the “control group”. These mussels were placed in a 200 liter seawater tank (Figure 2.6a)where they remained for the duration of the experiment. The water in this tank wascontinually replenished with fresh seawater which was pumped in from the ocean. Thesecond group, the “experimental group”, was placed in a wire cage and submersed in a 15liter seawater tank (Figure 2.6b). The water in this tank was not replenished and initiallycontained 45 tCi[3H]-MMA. Compressed air was bubbled through both of the tanks.The sea water in the smaller tank originated from the same source as used for the controltank. The mussels from the “experimental group” were left in the static tank for 3 daysbefore being transferred into the larger tank which also contained the “control group”.After 4 additional days, the “experimental group” was transferred back into the smallertank. This cycle of exposure followed by washing was continued for a total of 5 weeks.The water in the static tank was changed after 3 weeks and replaced with fresh sea waterwhich also contained 45 .tCi[3H1-MMA. The water was replenished so that the wasteproducts of the mussels would not accumulate to unhealthy levels for the mussels. At theend of the experiment, the mussels were left in the large tank for one day in order to washaway any radiolabeled compounds that were loosely attached to the surfaces of the musselshells and flesh.2.2.7 Isolation of the13H1-labeled arsenicals in the mussel fleshThe mussels from the “experimental group” (75) were shucked and the byssalthreads, shells and flesh were isolated and pooled. Each pool was washed with deionizedwater and then was set aside for subsequent total arsenic determinations. This processwas repeated for the “control group”.The following procedure for isolating arsenicals in the mussel flesh was27A)B)seawater from oceanFigure 2.6 A) 200 liter seawater tank with continually circulating seawater; B) 15 literexperimental tank containing[3HJ-MMA C) “control group” of mussels;D) wire cage containing “experimental group” of mussels; E) aerationstones and tubing.E)seawater to ocean28developed by M. Dodd46 and is summarized in Figure 2.7. Some flesh (5 g) from the“experimental group” was set aside for total arsenic determinations and the remaining flesh(65 g) was homogenized and extracted with MeOH (2.5 ml MeOH per gram of flesh).The resulting slurry was mechanically shaken for 2 days and then filtered to separate thesoft brown residue from the extract. Another MeOH extraction was performed on theresidue and both of the MeOH extracts were combined. The MeOH was evaporated(rotary), and the black-gummy residue was redissolved in a minimum amount ofwater (25ml). A small amount of this solution (200 j.tl) was set aside and counted. The remainingsolution was repeatedly extracted with 25 ml portions of diethyl ether until the etherextracts were colourless. The pooled ether extracts were also counted. The ether extractswere not further examined. The water fraction was evaporated (rotary) to dryness and theresulting residue was redissolved in about 10 ml of MeOH. This solution was applied to aSephadex LH-20 column (1.5 x 2.5 cm) and eluted with MeOH (450 ml). Fractions (10ml) were collected and counted by using the scintillation counter (100 .d samples). The[3Hj containing compounds eluted within one band and the appropriate fractions werecombined and evaporated (rotary) to dryness. The residue was redissolved in 10 ml ofwater and applied to a Dowex 50Wx8 cation exchange column (1.5 x 20 cm). (The resinin the [Hj form was prepared as follows: wash with 2 M NaOH (3 times), wash withdeionized water until neutral, wash with 2 M HCI (3 times) and then with deionized wateruntil neutral.) The[3H]-labeled compounds were eluted from the Dowex column by using100 ml H20 (fractions 1-9), 200 ml of 5% NH4O (fractions 10-30), 50 ml H20(fractions 3 1-35), and 150 ml 2 M HCI(fractions 36-50). Fractions (10 ml) were collectedand counted (100 il samples). The fractions containing[3H1-labeled compounds whicheluted in the NH4O fraction were combined and concentrated on the rotary-evaporator.29The fractions containing[3H]-labeled compounds which eluted in the HCI fraction weretreated in a similiar fashion. The fractions containing[311J-labeled compounds whicheluted with H20were not further analysed because these fractions were not expected tocontain either arsenobetaine or tetramethylarsonium ion.HomogenizeMussel—— MeOH extractConcentrateEther/H20partition -4 H20 (suspend)H20 layer- SephadexLH2ODowex5OWx8H20 NH3 HCIFigure 2.7 Schematic representation of the procedures used for the extraction ofarsenic compounds from mussels and for the purification of the extract.302.2.8 Identification of the[3HJ-labeled compound in the mussel flesh which elutedfrom the Dowex column in the NU4OH fractionThe residue in the N}L1OH fraction was redissolved in a minimum amount ofH20- (2 ml). A small aliquot was then applied via a drawn out capillary tube to a TLC plate (3x 20 cm strip of silica gel). Two different solvent systems were used as eluents. Solventsystem #1 was composed of ethanol-acetic acid-water (65:1:34) and solvent system #2was composed of acetonitrile-acetic acid-water (65:1:34). When the solvent had risenabout 16 cm the plates were removed from the solvent chamber and dried. The driedplates were cut into 16 segments (1 x 3cm) which were placed into 7 ml scintillation vialscontaining 5 ml of scintillation fluid. The vials were counted to determine which fractioncontained the radiolabeled compound(s).The[3H1-labeled compound(s) in the NH4O fraction were also examined byusing HPLC. A Waters Protein Pak DEAE anion exchange column was used and thesample was eluted by using 5mM sodium acetate (pH = 6.67) as the eluent at a flow rateof 1 ml per minute. Fractions were collected every minute and counted (100 i.l samples).2.2.9 Analysis of the[3H]-labeled compounds in the mussel flesh which eluted fromthe Dowex column in the HCI fractionThe compound(s) in the HC1 fraction were examined by TLC, using solvent system#1, as described in Section 2.2.8.2.2.10 Determination of the total arsenic concentration in the mussel flesh, byssalthreads and shellsThe determinations of the total arsenic levels followed the procedures developed31by Dodd.46 The flesh and the byssal threads for both the “experimental group” and the“control group” were decomposed by using a solution of 69% nitric acid (3 ml), 98%sulphuric acid (1 ml) and 30% hydrogen peroxide (3 ml). This mixture was added toapproximately 1gm of accurately weighed sample in a 250 ml round bottom flask. Theflask was fitted with a specially designed stopper and air condensor containing a diffusionfI.rnnel as shown in Figure 2.8. The reaction mixture was heated to a gentle reflux for 3hours and then allowed to cool. The digested sample was transferred into a 100 mlvolumetric flask and made up to the mark with deionized water.The shells were washed with deionized water and ground to a fine powder byusing a mortar and pestle. Hydrochloric acid (2M) was added to the powder. For everygram of shell 15 ml of acid was used. After an hour the resulting suspension was filteredand the filtrate was transferred to a 100 ml volumetric flask and made up to the mark withdeionized water. The total arsenic was determined in the samples by using HGAAspectroscopy. The digestate for both the “control group” and the “experimental group”was also counted. All digestions described above were performed in duplicate.32Figure 2.8 Wet ashing apparatus; (A) Teflon cylindrical plugs; (B) Teflon difflisionfunnels; (C) Teflon stopper with capillary; (D) 250 ml round bottom flask.All joints are 14.5/23. All dimensions are mm.LD.fl332.3 RESULTS AND DISCUSSION2.3.1 The uptake ofJ3111-Jabeled arsenicals from the seawaterThe mussels in the “experimental group” were exposed to seawater which initiallycontained[3H1-MMA for a five week cycle of exposure/washing. The mussels in the“control group” were not exposed to any[3H]-labeled compounds. At the end of theexperiment the mussels in the “experimental group” appeared to be as healthy as themussels in the “control group”, despite exposure to arsenic concentrations well in excessof the normal environmental levels. This statement is made after observing that the twogroups were not visually different and the effort needed to pry open the shells was aboutthe same. If a mussel is unhealthy the shell will open very easily and the flesh will notadhere to the inside of the shell very well. There were no casualties in either group duringthe five weeks.2.3.2 Purification of the[311]-labeled arsenicals in the mussel flesh of theexperimental groupThe[3H]-labeled compounds were extracted with MeOH from the homogenizedmussel flesh of the “experimental group” as outlined in Section 2.2.6. The diethyl etherextraction withdrew approximately 10% of the[3H]-labeled compounds from the MeOHflesh extracts. This indicates that some of the[3H]-Iabeled MMA has beenbiotransformed into more complex arsenicals which may include the arsenic containinglipids 7a and 7b. The water soluble[3Hj-Iabeled compounds were added to a Sephadexcolumn, eluted by using water, and the fractions were counted to yield the chromatogramin Figure 2.9. The[3H]-labeled compounds eluted from the column in one band and34IIFigure 2.9 The elution profile from a Sephadex LH-20 column for the water soluble[3H]-labeled compounds which were MeOH extracted from the mussels.FRACTION NUMBER351 28 37FRACTION NUMBER0£146Figure 2.10 The elution profile from the Dowex column for the[3H]-labeledcompounds which eluted in fraction 8 to 21 from the Sephadex column.36fractions 8 to 21 were combined. The combined fractions were concentrated, applied to aDowex column, and eluted by using H20, 5% NH4O and 2M HC1 to yield thechromatogram in Figure 2.10. A number of different[3H]-labeled compounds seem to bepresent with the bulk of the radioactivity being eluted in both the H20and the NFL1OHfractions. There also seems to be a small amount of[3Hj-labeled compounds in the HC1fraction. Little is known about the arsenic containing compounds which elute off of thistype of column in the water fraction and there are no standards for Rf comparison, so the[311j-labeled compounds in the water fraction were not further investigated. Fractionnumbers 11 to 17 eluting with a retention volume of between 110 ml and 180 ml wereconcentrated and designated the “NH4O fraction” for future reference. Fractionnumbers 39 and 40 eluting with a retention volume between 390 ml and 410 ml wereconcentrated and designated “the HCI fraction” for future reference.2.3.3 Identification of[3H]-arsenobetaine in the NH4O fractionThe compound(s) which eluted from the Dowex column in fractions 11 to 17 (theNH4O fraction) were further examined by using TLC with two different solventsystems. The results of comparing the Rfs of arsenobetaine standard and the unknown aredisplayed in Table 2.3.Table 2.3 Rf values for the unknown from the “NH4O fraction.”Solvent System Rf Arsenobetaine Rf Unknown#1 0.52 ± 0.05 0.55 ± 0.05#2 0.43 ± 0.05 0.43 ± 0.0537These results show that the[3H]-labeled compound eluting in the “NH4Ofraction” has the same Rfvalues as does arsenobetaine. The arsenobetaine standard waschosen because it is the major arsenical found in most marine organisms and its elution inthe “NH4O fraction” is well documented.’0°Additional evidence for this identificationis provided by anion exchange HPLC. The arsenobetaine and DMA standards were elutedthrough the anion exchange column, Figure 2.11 a, by using 5 mM sodium acetate. Theunknown compound was eluted through the anion exchange column, Figure 2.1 Ib, byusing 5 mM sodium acetate. The unknown and the arsenobetaine have the same retentiontime under these conditions. This result confirms the identification of the unknowncompound in the “NH4O fraction” as arsenobetaine and eliminates DMA (the otherknown arsenical which elutes off of the Dowex column in the “NH4O fraction”) as apossibility.The exposure of mussels to[3H}-MMA and the subsequent identification of[3HJ-arsenobetaine in the mussels confirms that MMA is a possible precursor to arsenobetaine.This is suggested by a combination of the schemes in Sections 2.1.3 and 2.1.4. This resultis significant because it is the first time that radiolabeled arsenobetaine has been isolatedfrom a marine organism after the organism had been exposed to a simple radiolabeledarsenic compound. This is unlike the experiments described in Section 2.1.1 with crabs,fish and gastropods where[74As]-arsenate was investigated as the precursor toarsenobetaine. The success of the experiment described in this chapter may be due to thefact that[3Hj-MMA was added directly to natural seawater which contained all differenttypes of algae, bacteria, phytoplankton, diatoms and nutrients. Thus, the naturalconditions were duplicated as closely as possible, except for the exclusion of thesediments. The experiments which attempted to use[74As]-arsenate as the precursoreither fed the precursor to the marine organism directly or cultured one specific strain of380.09A)0.080.07’w 0.060z 0.05o 0.04C,)< 0.030.020.010B)0Figure 2.11LI I I I 1 I I I10 19FRACTION NUMBER1 2 3 4 5 6 7 8 91011121314151617181920FRACTION NUMBERHPLC chromatograms from the anion exchange column at pH 6.67; (A)arsenobetaine and dimethylarsinate on the anion exchange column at pH6.67, (B) the compound eluting in fractions 11 to 17 from the Dowexcolumn.39algae with the precursor. Thus, any complex interactions between various marineorganisms which may be necessary for this biotransformation to occur would not bepossible under these conditions. The experiment described in this chapter also had a muchlonger duration than the previous experiments and it is possible that this biotransformationis a relatively slow process. In the proposed scheme, Section 2.1.4, it is suggested that inorder for the C3-C4 bond in the arsenic containing riboside to be cleaved, anaerobicconditions are necessary. It was suggested that this cleavage may occur in the sediments,however, the experiment described in this chapter demonstrates that arsenobetaine can beformed in the absence of any sediments.2.3.4 Attempted identification of the[311]-containing arsenicals in the HCIfractionIdentification of the[3H]-labeled compound(s) in the “HCI fraction” washampered because of the relatively low level of radioactivity in this fraction. The twomajor arsenicals known to elute off of the Dowex in the “HCl fraction” are the cationictetramethylarsonium ion and arsenocholine. TLC analysis of the sample was inconclusiveas shown in Table 2.4. Lack of additional sample made further analysis impossible. Fromthe results it would be quite reasonable to suggest that the unknown[3H]-labeledcompound(s) eluting off of the Dowex column in the “HCl fraction” could have beeneither arsenocholine or tetramethylarsonium ion. Arsenocholine is postulated to be theimmediate precursor to arsenobetaine, (see Section 2.1.4) however, it has not been foundto occur naturally in mussels. Tetramethylarsonium ion is present in most types ofmussels.5640Table 2.4 The[3H]-containing compound(s) in the “HCl Fraction.”Solvent System Rf tetramethyl- Rf arsenocholine Rf Unknownarsonium ion#1 0.38 ± 0.05 0.32 ± 0.05 0.35 ± 0.052.3.5 Total arsenic concentrations in the mussel flesh, byssal threads and shells.The mussels were sacrificed after five weeks of intermittent exposure to the[3H)-labeled MMA and any other[3H]-labeled compounds that may have been produced by themicro-organisms present in the seawater. The “control group” and “experimental group”of mussels were shucked and the flesh, byssal threads and shells for each group werepooled. These three components were acid digested as outlined in Section 2.2.10. Theinitial arsenic concentration of[3H]-MIvIA that the mussels were exposed to was 2 ppm.The total arsenic concentrations in the mussel flesh, byssal threads, and shells of both the“experimental group” and the “control group” are summarized and displayed in Table 2.5.The natural arsenic concentration in the seawater that was being pumped into the tankswas determined by using HGAA spectroscopy to be 0.0054 ppm.Table 2.5 Arsenic concentration in the mussel flesh, byssal threads, and shells.As concentration (ppm) As concentration (ppm)“Control group” “Experimental group”Flesh 0.45 ± 0.04 0.87 ± 0.07Shells 0.24 ± 0.03 0.32 ± 0.04Byssal threads 4.20 ± 0.20 6.30 ± 0.3041The total arsenic concentration in each of the three compartments increasedsignificantly during the course of the exposure experiment. This result is expected as themussels were intermittently exposed to an arsenic concentration that was approximately400 times the natural level. The arsenic concentration in the byssal threads is muchgreater (10 times) than the concentration in the flesh or the shells. The excretion ofheavy metals into the byssal threads by the mussels is thought to be part of thedetoxification mechanism of the organism.105 The digestion procedure for both the fleshand the byssal threads would have decomposed any organoarsenicals which may have beenpresent, thus counting techniques were not employed. The procedure used to dissolve theshells would not have led to the complete degradation of the organoarsenical, in particularthe bond between the As and[3H1-methyl may not been cleaved, so this digestate wascounted to yield the results displayed in Table 2.6.Table 2.6 The radioactivity in the shells.“Control group” “Experimental group”Shells 22.15, 22.95 DPM 32.43, 31.26 DPMCompounds may be either adsorbed onto the shell from the surrounding media orincorporated into the shell by the mussel. The percentage increase in the arsenicconcentration in the shells corresponds almost exactly with the increase in radioactivity.This result suggests that the arsenicals which were incorporated into the shells during theexperiment were methylated. This finding is consistent with the finding of low levels of42methyl- and dimethyl- arsenicals in the shells of some costal bivalves ofBritish062.3.6 SummaryThe results described in this chapter show that feeding-type experiments can yieldvaluable infomation about the biotransformations of arsenic in the marine environment, inparticular that MMA is a precursor to arsenobetaine. As a cautionary note, thebiotransformation was observed in a situation where the arsenic levels were elevated toabout 400 times the natural concentration. This biotransformation occurs in the absenceof the strongly anaerobic conditions which are found in the sediments. These conditionswere thought to be necessary for the decomposition of the arsenosugars which have beenpostulated to be key intermediates in the scheme for arsenobetaine formation. Other[3Hj-labeled compounds were present in the mussels, but were not positively identified.The overall arsenic concentration increased in the mussel flesh, byssal threads and shellsduring the course of the experiment. Data were also collected which suggests that thearsenicals present in the shells were methylated.The results from this chapter do not, however, establish the source of thearsenobetaine. The biotransformation either is performed by the microscopic organisms(for the purposes of this thesis, the term microscopic organisms will refer to any organismwhich is not normally seen by the human eye) in the seawater, or by the musselsthemselves. Another possible scenario is a combination of the two possibilities. Clearly,these experiments should be modified in order to answer these questions (see Chapter 3).43CHAPTER 3THE BIOTRANSFORMATION OF MMA AND DMA INTO ARSENOBETAINEIN SEAWATER AND MUSSELS3.1 INTRODUCTION3.1.1 The origin of arsenobetaine in seawater and marine animalsArsenobetaine is the major form of arsenic in most marine animals. There are twopossible ways for marine animals to acquire arsenobetaine. Either it is accumulateddirectly from food and/or seawater or it is synthesized from other arsenicals within theanimal. Studies in this area have not been fruitful, partly because the analytical techniquesavailable for determining arsenic speciation in seawater are generally not suitable for thedetection of arsenobetaine. There have been no previous reports of the presence ofarsenobetaine in seawater, although it was estimated by Francesconi99that if it waspresent, its concentration would be less than 0.5 jig As L1.Francesconi has also examined the ability of marine animals to bioaccumulatearsenobetaine, arsenocholine, dimethylarsinylethanol, and dimethylarsinylacetic acid. Heshowed that both arsenobetaine and arsenocholine were readily taken up by the yelloweyemullet from raw beef which was fed to the fish.103 The raw beef had been injected withboth arsenobetaine and arsenocholine. When the same experiments were carried out withdimethylarsinylethanol and dimethylarsinylacetic acid in the raw beef there were noincreases in the arsenic concentration in the yelloweye mullet. Similar experimentsinvolving adding arsenobetaine directly to a tank of seawater which contained mussels (Medulis) showed that the mussels were unable to regulate their uptake of arsenobetainefrom the seawater. The amount of arsenobetaine in the mussels at the end of the44experiments was simply dependent upon the amount of arsenobetaine in the seawater andthe amount of time that the mussels were exposed to it.99 When trimethylarsine oxidewas added directly to a tank of seawater which contained mussels (M edulis), thetrimethylarsine oxide was not appreciably accumulated. In the experiments whichinvolved arsenobetaine accumulation, the marine animals did not metabolizearsenobetaine.The results from experiments which investigated the ability of marine animals tosynthesize arsenobetaine from algal arsenic are outlined in Chapter 2, Section 2.11.3.1.2 Scope of workThe objective of the experiments described in this chapter was to modify thefeeding-type experiments described in the previous chapter to answer the major questionswhich arose in the interpretation of the results. What is the source of the arsenobetaine inthe mussels? Is the biotransformation of MMA into arsenobetaine performed by themicroscopic organisms in the seawater, by the mussels, or by some combination of the twopossibilities? These questions were answered by investigating the quantities andspeciation of the[3Hj-labeled arsenicals during the course of the modified feeding-typeexperiments.In this experiment either[3Hj-MMA or[311j-DMA was added to seawater bothwith and without the presence of mussels. After exposure the[3H]-labeled arsenicals inthe mussel flesh and the seawater were identified or an attempt was made to identify them.453.2 EXPERIMENTAL3.2.1 InstrumentationThe instrumentation and techniques for scintillation counting, atomic absorptionspectrometry, and HPLC were identical with those described in Sections 2.2.1, 2.2.2, and2.2.3.3.2.2 Chemicals and reagentsThe chemicals and reagents used in this chapter are identical to those described inSection 2.2.5.3.2.3 The uptake and conversion of13H1-MMA and[3H]-DMA toi311]-arsenobetaine in seawater and musselsApproximately 75 mussels (M edulis), ranging in shell length from 2.5 to 5.0 cm,were collected from the beach surrounding Point Grey in Vancouver B.C. They wereplaced in a wire cage which was submerged in a 15 liter seawater tank. Another 15 litertank was set up as the control. The seawater sampled from the tank containing musselswill be referred to as the “experimental seawater” and the seawater sampled from the othertank will be referred to as the “control seawater”. The water in these tanks was notreplenished and initially contained either[3HJ-MMA or[3HJ-DMA at an activity level of45 i.Ci. The mussels were left in the 15 liter tank for 4 days before being transferred intoa third tank which contained 200 liters of seawater. The water in this tank was continuallyreplenished with fresh seawater which was pumped in from the ocean. The seawater in allthree tanks originated from the same source and aeration was achieved by the use ofcompressed air. The same tanks shown in Figure 2.6 were used with the addition of a46second static tank which held only the control seawater. On days 4 and 7 the“experimental seawater”, “control seawater”, and mussels were sampled. This samplingprocedure involved the collection of200 ml of both the experimental and control seawateralong with 10 medium sized mussels.3.2.4 Isolation and identification of the13H1-labeled arsenicals in the seawater andthe mussel fleshThe “experimental seawater” samples (200 ml) were filtered through a hydrophilicmembrane filter (0.45 p.m pore size). The pH of the filtrate was adjusted to 4 by adding afew drops of concentrated HC1. This solution was counted (200 p.1) and extracted withphenol (2 x 50 ml). The combined phenol extracts were diluted with diethyl ether (400ml) and the water soluble compounds were extracted with deionized water (2 x 50 ml).The aqueous extracts were combined, counted (200 p.1), evaporated (rotary) to drynessand redissolved in deionized water (10 ml). This solution was applied to a cationexchange column (Dowex 50Wx8, 1.5cm x 20cm in the [Hj form). The[3H]-labeledcompounds eluted from the column by using deionized water (70 ml) and 5% ammoniumhydroxide (200 ml). These conditions were sufficient to elute all of the[3H]-activity.Fractions were collected (20 ml) and counted (600 p.1 samples). The H]-containingfractions were bulked, evaporated (rotary) to dryness, and redissolved in a minimumamount of deionized water (1 ml). Aliquots (100 p.1) of these solutions were examined byusing two different types ofHPLC columns. The first, a Protein Pak DEAE column wasrun by using 5 mM sodium acetate at pH 4.0 and pH 6.67 as the eluant. The second, aProtein Pak 300SW column was run by using 5 mM sodium acetate at pH 6.67 as theeluant. The flow rate was set at 1 ml. per minute and fractions were collected every minute47and counted (100 p.1 samples). Standard solutions of MMA, DMA and arsenobetaine indeionized water were chromatographed on both the cation exchange and HPLC columns,fractions were detected by using GFAA spectroscopy. The “control seawater” wastreated in an identical fashion.The mussels were washed with deionized water and the shells were removed fromthe flesh which was then homogenized. The resulting slurry was made up to 200 ml byusing deionized water. This slurry was treated in an identical fashion to the 200 mlseawater filtrate.3.2.5 The diversity of[3H]-labeled compounds in the seawaterThe experimental conditions in this section were virtually identical with thosedescribed in Section 3.2.3, except for the following: only[3H]-MMA was used as thestarting substrate, the duration of this experiment was 10 days and the mussels weretransferred from the large tank where the seawater was circulating to the smaller tankwhere the seawater was not circulated on day 7. Seawater samples were collected on days4, 7 and 10 from the smaller tank which contained the[3H]-labeled compounds.3.2.6 Examination of the13H1-Iabeled compounds in the seawater from theexperiment described in Section 3.2.5The seawater was subjected to the set of extractions which were described inSection 3.2.4 and the extracted compounds were applied directly to a size exclusionHPLC (HPLC Protein Pak 3 OOSW) column. The[3Hj-labeled compounds were eluted byusing 5mM sodium acetate at a flow rate of imi per minute. Fractions were collectedevery minute and counted (100 p.1 samples).483.3 RESULTS AND DISCUSSION3.3.1 Experimental seawater after exposure to[311]-MMA and[3H]-DMA[3HJ-MMA or[3H]-DMA was added to seawater which contained mussels(M edulis). After four days the mussels were transferred into another tank. The seawaterfrom the tank which contained the mussels was examined on days 4 and 7. The seawaterwas filtered, counted, extracted with phenol, back extracted with deionized water, andthen counted again. The chromatographic results and identification of the[311]-labeledcompounds phenol extracted from the seawater are described below.It is important to note that there is no possibility of proton exchange in situ for thecompounds which are being investigated. Any casual exchange of the label betweencompounds would require a cleavage of the methyl-arsenic bond. Also, the various[3H]-labeled compounds in the seawater will all have different extraction efficiencies intophenol, therefore, the ratio of compounds in the extracts will be different from the actualratio of compounds in the seawater. Both MMA and DMA have extraction efficiencies ofabout 10%, whereas the extraction efficiency for arsenobetaine is closer to 100%.a)[3H]-MMA:The cation and anion exchange chromatograms of the standards and of the[3H]-containing compounds extracted from the experimental seawater on days 4 and 7 aredisplayed in Figures 3.1, 3.2, 3.3, and 3.4.The compound eluting in fraction 3 from the cation exchange column (Figure 3.2)was identified as MMA, on the basis of its retention time which was identical with that ofthe standard (Figure 3.1). This identification was confirmed by the chromatograms inFigures 3.3 and 3.4 for the elution profiles from the anion exchange (pH 4.0) and sizeexclusion HPLC columns, respectively.49The compound eluting in fraction 7 from the cation exchange column (Figure 3.2)was identified as arsenobetaine on the basis of its chromatographic coordinates whichwere identical with that of the standard. This identification was confirmed by thechromatograms in Figures 3.5 and 3.6 for the elution profiles from the anion exchange(pH 6.67) and size exclusion HPLC columns, respectively.wC)z0Cl)Figure 3.1 The cation exchange chromatograms for the standards of MMA, DMA,and arsenobetaine.FRACTION NUMBER500C12001000•800600400-200-0•1 2A)B)Figure 3.24 5 6 7FRACTION NUMBER8 9 110456 7FRACTION NUMBERThe cation exchange chromatograms of the phenol extract from the“experimental seawater” on days 4 (A) and 7(B) after[314}-MMAexposure.51A)B)350300250200150100500Figure 3.31 2 3 4 5 6 7 8 9 1011121314151617181920FRACTION NUMBERHPLC chromatograms; (A) arsenobetaine, MMA, and DMA on theanion exchange column at pH 4, (B) the compound eluting in fraction 3(Figure 3.2 (B)) on the anion exchange column at pH 4.wC)zcc0C,)a:i< 0.1400-FRACTION NUMBER52A)B)000FRACTION NUMBERFigure 3.4 HPLC chromatograms; (A) MMA on the size exclusion column and (B)of the compound eluting in fraction 3 (Figure 3.2 (B)) on the size exclusioncolumn.wC)z0Od0.1FRACTION NUMBER53A)0.090.080.07w 0.06C-)z< 0.050 0.04C,)< 0.030.020.01I IF1 10 19FRACTION NUMBERB)350-300-250-200-0150-100-s.1 2 3 4 5 6 7 8 91011121314151617181920FRACTION NUMBERFigure 3.5 HPLC chromatograms; (A) arsenobetaine and DMA on the anionexchange column at pH 6.67, (B) the compound eluting in fraction 7(Figure 3.2 (B)) on the anion exchange column at pH 6.67.54A)B)a0Figure 3.6 HPLC chromatograms; (A) arsenobetaine on the size exclusion column,and (B) of the compound eluting in fraction 7 (Figure 3.2 (B)) on the sizeexclusion column.wC)zEn00.1FRACTION NUMBER10 19FRACTION NUMBER55These results describe for the first time the isolation of arsenobetaine fromseawater. The isolation of arsenobetaine from the present experiments may have beenfacilitated by the use of elevated arsenic concentrations and by the use of a radioactivelabel to allow easier detection.The chromatograms in Figure 3.2 show a change in the relative amounts ofarsenobetaine and MIvIA on days 4 and 7. The amount of arsenobetaine increased in theabsence of mussels which indicates that it is being bio-synthesized, at least in part, bymicroscopic organisms which are present in seawater.b)[3H]-DMA:The cation exchange chromatograms of the[3H]-containing compounds extractedfrom the “experimental seawater” are displayed in Figure 3.7. The most notable feature ofthese chromatograms is the low level of activity, which is barely above the background.Arsenobetaine is tentatively identified in fraction 7 (Figure 3.7) by the chromatographictechnique described above. The elution profile from the anion exchange (pH 6.67) HPLCcolumn for the compound(s) eluting in fraction 7 (Figure 3.7 (B)) from the cationexchange column is displayed in Figure 3.8.56A)B)C0ci01FRACTION NUMBER2 3 4 5 6 7 8 9FRACTION NUMBERFigure 3.7 The cation exchange chromatograms for the experimental seawater on days4 (A) and 7 (B) after[3H)-DMA exposure.5710080600:: rdIbiuibiiHiiii1 2 3 4 5 6 7 8 91011121314151617181920FRACTION NUMBERFigure 3.8 The IIPLC chromatogram for the compound eluting in fraction 7 (Figure3.7 (A)) on the anion exchange column at pH 6.67.3.3.2 Controi seawater after exposure to[311]-MMA and13H1-DMAThe seawater from the control 15 liter tank was filtered, counted, extracted withphenol, back extracted with deionized water, and then counted again as described above.The chromatographic results and identification of the[3H]-labeled compounds extractedfrom the seawater are described below. Either[3H]-MMA or[3H]-DMA was added tothe seawater at the beginning of the experiment.a)[3H]-MMA:The cation and anion exchange chromatograms of the[3H]-containing compoundsextracted from the “control seawater” are displayed in Figures 3.9 and 3.10.58Figure 3.9 The cation exchange chromatograms for the phenol extract from the“control seawater” on days 4 (A) and 7 (B) after[3H]-MMA exposure.A)0030004 5 67FRACTION NUMBER2500200015001000500-0-12 345678910FRACTION NUMBER59800700600500400300200io:-1 2 3 4 5 6 7 8 91011121314151617181920FRACTION NUMBERFigure 3.10 The HPLC chromatogram for the compound eluting in fraction 7 (Figure3.9 (B)) on the anion exchange column at pH 6.67.The compounds eluting in fractions 3 and 7 from the cation exchange column(Figure 3.9) were identified as MMA and arsenobetaine by using the chromatographictechniques described above. Arsenobetaine is confirmed by the chromatogram in Figure3.10 for the elution profile from the anion exchange HPLC column at pH 6.67.These results confirm that MMA can be converted to arsenobetaine in naturalseawater. Mussels were never present in the tank so the transformation must beaccomplished by organisms at lower trophic levels. The ratio of arsenobetaine to MMApresent in the control (Figure 3.9) is much less than that found in the “experimentalseawater” (Figure 3.2). This could mean that arsenobetaine is being synthesized by60mussels and/or other microscopic organisms that are symbiotic or otherwise present inseawater with the mussels. It is impossible to distinguish between the contributions of theorganisms associated with the mussels and those of the mussels in the current set ofexperiments.b)[3H]-DMA:The cation and anion exchange chromatograms of the[3Hj-containing compoundsextracted from the “control seawater” are displayed in Figures 3.11 and 3.12.The compound eluting in fraction 7 from the cation exchange column (Figure 3.11)is identified as arsenobetaine by using the chromatographic techniques described above.Arsenobetaine is confirmed by the chromatogram in Figure 3.12 for the elution profilefrom the anion exchange HPLC column at pH 6.67.It seems that DMA is more easily biotransformed to arsenobetaine than MMA.This conclusion is supported by the work in Chapters 4 and 5 which indicates that DMApasses more freely through model cell membranes than MMA. Thus, it would be morereadily available for biotransformation.61A)1600140012001000800600400-200-—B)50O—450-400350-300-250-200-1 50-10050-— I- -T1 2 3 4 5 6 7 8 9 10FRACTION NUMBERFigure 3.11 The cation exchange chromatograms for the phenol extract from the“control seawater” on days 4 (A) and 7 (B) after[3H]-DMA exposure.1 2 3 4 5 67 8 9 110FRACTION NUMBER62800-700600500400300200100I I1 2 3 4 5 6 7 8 91011121314151617181920FRACTION NUMBERFigure 3.12 The HPLC chromatogram for the compound eluting in fraction 7 (Figure3.12 (B)) on the anion exchange column at pH 6.67.3.3.3 Mussel Flesh after exposure to[3111-MMA and[3Hj-DMAAfter exposure of the mussels to the labeled arsenicals the flesh was homogenized,the[3H)-containing compounds were extracted from the flesh with phenol and then backextracted into water.a)[3H]-MMA:The cation exchange and anion exchange chromatograms of the[3}Ij-containingcompounds phenol extracted from the mussel flesh after 4 days exposure to the label and63A)B)1:80-UFRACTION NUMBER1 2 3 4 5 6 7 8 9 10FRACTION NUMBERFigure 3.13 The cation exchange chromatograms for the phenol extracted compoundsfrom the mussel flesh on days 4 (A) and 7 (B) after[3HJ-MMA exposure.64after the ftill 7 day cycle (4 days exposure, 3 days wash) are displayed in Figures 3.13 and3.14.The compound eluting in fraction 7 from the cation exchange column (Figure 3.13)is identified as arsenobetaine by using the chromatographic techniques described above.The chromatographic behaviour of the compound in fraction 2 of Figure 3.13 doesnot match that of any of the available standards. Also, the elution profile for thiscompound from the anion exchange HPLC column shown in Figure 3.14, does notcompare with that of MMA standard shown in Figure 3.3. Its chromatographic behaviourindicates that it is neutral, as it elutes in the void volume from both the cation and anionexchange columns. Arsenic is present in the fraction (by using GFAA) and its possiblethat the arsenical is a neutral arsenosugar such as 2’,3’-dihydroxypropyl 5-deoxy-5-dimethylarsinyiriboside 6a.250-2001 50010050 1 H1s1u123456 7 8 91011121314151617181920FRACTION NUMBERFigure 3.14 The HPLC chromatogram for the compound eluting in fractions 2 and 3(Figure 3.13 (B)) on the anion exchange column at pH 4.65The two chromatograms in Figure 3.13 are quite similar in that the ratio ofarsenobetaine to the unknown is almost the same. Between days 4 and 7 the mussels wereexposed to continuously circulating seawater (from the ocean) and any[3Hj-labeledcompounds that were not bound to the mussels would be quickly washed away. Thus, thechromatograms indicate that within the experimental uncertainty the mussels are not biosynthesizing arsenobetaine from any of the compounds they have accumulated while beingexposed to labeled MMA. This finding supports the hypothesis that arsenobetaine inmussels is accumulated from the surroundings.b)[311]-DMA:The flesh of the mussels exposed to labeled DMA was examined in a similarmanner. The compound eluting in fraction 7 from the cation exchange column (Figure3.15) is identified as arsenobetaine by means of the chromatographic techniques describedabove.The compound in fraction 2 appeared to be the same unidentified arsenical isolatedfrom the M1’vIA exposure experiment. Again, as in Figure 3.13 the two cation exchangechromatograms (Figure 3.15) are quite similar. This supports the idea that musselsselectively accumulate arsenicals, as was shown by Francesconi in controlled uptakeexperiments where mussels readily bio-accumulated arsenobetaine and arsenocholine, butdo not bio-accumulate trimethylarsine oxide or 2-dimethylarsinylethanol.99These results indicate that either MMA or DMA can act as a precursor toarsenobetaine and are part of the same bio-synthetic pathway.66A)B)a0a0FRACTION NUMBER4 5 6 7FRACTION NUMBERFigure 3.15 The cation exchange chromatograms for the phenol extracted compoundsfrom the mussel flesh on days 4 (A) and 7 (B) after[3HJ-DMA exposure.673.3.4 The extraction efficiency of[3H]-labeled compounds which were extractedfrom seawater and mussel samplesEach of the samples was counted prior to and after the phenol extractions in orderto determine the percentage of[3111-labeled compounds which were extracted from thematrix by the phenol. These results are summarized in Table 3.1.Table 3.1 Percentage of[3H]-labeled compounds which were extracted from theseawater and mussel samples.“experimental “control musselsseawater” seawater”MMA starting 10% 10% 80%DMA starting 5% 40% 85%The extraction efficiency of the phenol extraction procedure (Section 3.2.4) forboth MIvL& and DMA was experimentally determined to be about 10%. The phenolextractions were designed by Cannon et alto extract arsenobetaine from a complexsample matrix.42 The extraction efficiency of the phenol extraction proceedure forarsenobetaine was experimentally determined to be close to 100%. The results from Table3.1 show that there is a much higher percentage of[3H]-labeled compounds extractedfrom the mussels than from the seawater. This indicates that the composition of[3H]-labeled arsenicals in the mussels is different than in the water. This difference supports thenotion that mussels selectively accumulate organoarsenic compounds from seawater,rather than randomly accumulating compounds at the percentages at which they are found68in the seawater (as was discussed in Section 3.3.3). When DMA, rather than MMA wasused as the starting substrate in the “control seawater” there is a greater percentage of[3Hj-labeled compounds extracted from the seawater. This greater percentage indicatesthat more biotransformations to arsenobetaine and compounds similar in structure toarsenobetaine have occured. Such a finding supports the earlier results which suggest thatDMA is more readily biotransformed into arsenobetaine than is MMA (discussed inSection 3.3.2). The low percentage of[3H1-labeled compounds extracted from the“experimental seawater” when[3Hj-DMA was the starting substrate agrees with theearlier results (Section 3.3. ib) which showed that level of radioactivity in the extractsfrom the seawater was quite low. This tends to suggest that the rate of conversion of the[3H]-DMA to[3H]-arsenobetaine by the microscopic organisms in the seawater followedby the uptake of the{H]-arsenobetaine by the mussels must occur rapidly within the timescale of the experiment. This is consistent with the observed depletion of the[3Hj-label inthe seawater.3.3.5 Examination of the[3H]-Jabeled compounds in the seawater after theexperiment described in Section 3.2.5The seawater collected on days 4, 7 and 10 was filtered and the filtrate wassubjected to the phenol extraction proceedure (outlined in Section 3.2.6). Approximately10% of the[3111-labeled compounds in the seawater were extracted by the phenolextractions. The extracted compounds were back extracted into deionized water andevaporated (rotary) to dryness. This was redissolved in 1 ml of deionized water andapplied to a size exclusion NPLC column. The compounds were eluted by using 5 mMammonium acetate and the resulting chromatograms are displayed in Figures 3.16 A, B69and C for days 4, 7 and 10, respectively. The chromatogram in Figure 3.16 A from day 4shows that the majority of the extracted[3Hj-labeled compounds are eluted betweenfractions #12 and #16. This indicates that the[3H]-label is distributed in the seawateramong compounds of a similar size. The size exclusion chromatograms for the standardsMMA, DMA,, and arsenobetaine are shown in Figures 3.4 A, 3.17 and 3.6 A, respectively.Each of these three standards elute with approximately the same retention volume. Thechromatogram in Figure 3.16 B from day 7 is quite different than the chromatogramproduced on day 4 because most of the extracted[3H]-labeled compounds are elutedbetween fractions #11 and #30 showing that the extracted[3H]-label has beenredistributed amongst compounds of different sizes. This redistribution of the labeloccurred during the time when mussels were not present in the seawater. Therefore, aswas shown in the previous experiments involving the “control seawater”, the small, non-circulating tank contains a wide variety of microscopic organisms which are capable of thebiotransformation of arsenic. The mussels were placed back into the seawater betweendays 7 and 10 and the resulting chromatogram for day 10 is shown in Figure 3.16 C. Thischromatogram resembles the chromatogram from day 4 more closely than it resembles thechromatogram from day 7. The amount of extracted[3Hj-labeled compounds elutingbetween fractions #19 and #30 has decreased in the chromatogram for day 10 whencompared to the chromatogram for day 7. This suggests that the mussels may bebioaccumulating or biosynthesizing the compounds found in the latter chromatographicfractions.This particular study is inconclusive, but it does accentuate the fact that the arseniccycling in the ocean is based upon complex interactions between microscopic andmacroscopic organisms.wCDPMDPMCA)10CA)4010010010010o000000000000000000000000000IIIIIIIIIIIIICD2o o — CD CD CD 5no-______—Z____Z Cw___________________wmm,Ar4r4CDrr4rw4Wr4713002502000- 150100500C)IiiIhIhihhI I I I I I I I 1 I I I II I I I I I I I I1 10 19 28FRACTION NUMBERThe size exclusion HPLC chromatogram for the “experimental seawater”on days 4 (A), 7 (B), and 10 (C).Figure 3.16w 0.0z0C’)1 10 19FRACTION NUMBERFigure 3.17 The size exclusion HPLC chromatograms for the DMA standard.723.3.6 SummaryThe results from this chapter indicate that both MMA and DMA may bebiotransformed to arsenobetaine in the marine environment. In particular, it was shownthat naturally occuring microscopic organisms are capable of performing thisbiotransformation. Within the uncertainty of the experiments described in this chapter itdoes not appear that the mussels are capable ofbiosynthesizing arsenobetaine. Thus,mussels appear to accumulate their arsenobetaine content via uptake from the watercolumn. The results also show that mussels accumulate compounds selectively, as theybioaccumulate both arsenobetaine and an unknown compound, while the startingsubstrates are not bioaccumulated by the mussels within the detection limits of theexperiments.DMA is biotransformed more readily to arsenobetaine than is MMA. This resultindicates that DMA is either more direct link to arsenobetaine in the biochemical pathway,or that DMA is more rapidly bioaccumulated by the microscopic organisms in seawaterthan is MMA. The results of work described in Chapters 4 and 5 predict that DMA willbe more rapidly bioaccumulated by microscopic organisms than MMA.Future work may involve trying to isolate the microscopic organisms in seawaterwhich perform the biotransformations described in this chapter and the previous chapter.73CHAPTER 4THE USE OF LIPOSOMES IN PREDICTING THE BIOMOBILITY ANDBIOACCUMULATION OF MMA AND DMA4.1 INTRODUCTION4.1.1 How do molecules get across membranes?The translocation of matter across biological membranes is a fundamental physicochemical processes in living systems. The two major components of biological membranesare phospholipids and proteins (Figure 4.1). The proteins are inserted into phospholipidswhich are orientated in a bilayer.145 These bilayers form a continuous sheet around cellsand cellular organelles. For a molecule to enter or leave the cell or organelle it must crossthis barrier.Molecules may cross this barrier either by active transport or by passive diffusion.Active transport typically requires metabolic energy and is defined as the movement of ametabolite or an inorganic ion across a membrane usually in the direction that is againstthe concentration gradient. Passive (simple) diffusion occurs when molecules and ionscross either the lipid region of the barrier or flow through the aqueous channels formed bythe proteins, in a nonspecific way. This movement is driven solely by the difference in thechemical or electrochemical potential of the species on the two sides of the membrane.The lipid region is a hydrophobic environment. Thus, if the permeant is to diffusethrough this region of the membrane all of the bonds that the permeant has made with theaqueous medium surrounding the membrane must be broken. It is not thermodynamicallyfavorable for a charged molecule to enter into the hydrophobic environment of the innerbilayer.144 If a molecule does succeed in entering the hydrophobic region of the74Figure 4.1 The Fluid-Mosaic model of the cell membrane. The arrows represent thedirections of the lateral diffusion of the proteins.membrane by passing through the tightly packed head groups which present a significantbarrier to diffusion, then it will pass through the flexible hydrocarbon tails relatively easilyas this region has a much lower barrier to intramembrane movement.’46There are two possible mechanisms for the diffusion of molecules across the cellmembrane in which the permeant does not have to enter the hydrophobic core of thebilayer. Membranes are typically in a fluid state which allows small molecules to difihseinto and out of the vacancies (defects) in the outer bilayer (Figure 4.2a), thus moving froman aqueous environment into the apolar interior of the membrane. If a vacancy in the75lattice of the inner leaflet becomes available the permeant can then diffuse across themembrane. Membrane pores could also be formed by either the micellar arrangement ofthe phospholipid bilayers (Figure 4.2b) or by protein channels (Figure 4.2c) within themembrane. Thus, the permeant could pass through the bilayer without being impeded bythe hydrophobic region.b) c)JJUUFigure 4.2 Mechanisms of permeation through the bilayer without entry into thehydrophobic region.a) SIfJuSc5S764.1.2 Structures of main phospholipids found in cell membranesThe structures of the six major phospholipids found in cell membranes aredisplayed in Figure 4.3. For the purposes of this thesis, phosphatidyicholine was the onlyphospholipid used. Phosphatidyicholine is the most common lipid found in eukaryoticplasma membranes and is a zwitterion composed of a glycerol-phosphate ester with a0IO HC—O-C’RI I IiHO—P—0043 0OHPA00 H-O—C—RS ‘ II I U,C_CN_cH3_O r_O Q1: 0,Io OHPS0o )Ic!-O—C—RH I IIH.C—O—P—O—O4 0R—C-O—O1 OHR—C.O—CflIt0DPG0H03HC01Ro 11C’O—C—RI I UoroaIz0CuOHP10H C — 0 —O HC’O-C—R• II I U0OHPE0I0 HC—O—C—RH I H0013 OHPCFigure 4.3 Structures of the major phospholipids found in cell membranes; PA,phosphatidic acid; PS, phosphatidyl serine; PC, phosphatidyl choline;PE phosphatidylethanolamine; P1, phosphatidylinositol; DPG,diphosphatidylglycerol. R is the side chain (Figure 4.4).77choline head group and two acyl chains.’3’ The fatty acid side chains (Figure 4.4) of eggderived phosphatidylcholine are palmitic acid (32%), stearic acid (15%), oleic acid (31%),and linoleic acid (16%).132 In addition, there are other fatty acids present in traceamounts in egg phosphatidylcholine.a) CH3(CH2)14C02Hb) CH3(CH2)16C02Hc) CH3 (CH2)7CHCH(CH2)7C02Hd) CH3 (CH2)4(CH=CHCH2)2(CH2)6C02HFigure 4.4 The structure of the side chains of egg phosphatidyl choline; (a) palmiticacid, (b) stearic acid, (c) oleic acid, and (d) linoleic acid.4.1.3 Structures of hydrated phospholipidsWhen the dry phospholipids are hydrated a variety of lipid structures are formed.These structures are displayed in Figure 4.5. The structure most important to this thesis isthe liposome. Liposomes are single walled (unilamellar) or multiwalled (multilamellar)concentric shells in a spherical structure. A small volume of water is separated by thebilayer from the bulk aqueous solution (as shown in Figure 4.5c). The head groups areimmersed in the water; the hydrocarbon chains, being hydrophobic, point away from thewater to the center of the bilayer. This is energetically the most favorable arrangement, asthe water molecules are at a far lower energy when they are surrounded by other water78IitIABI)Structures ofphospholipids which are formed upon hydration;A) a monolayer formed at an air-liquid interface; B) a micelle; C) anexample of a liposome, (see Section 4.1.6); D) a black membrane,a planar bilayer, that separates two aqueous phases.O.CpFigure 4.579molecules with which they can hydrogen-bond, than when this bonding is interupted byhydrocarbon chains. Thus, water will be excluded from the region of the hydrophobic,hydrocarbon chains. Likewise, the hydrocarbon chains will line up with one another toform bilayers, which is the most thermodynamically favorable configuration.4.1.4 The mobility of the lipid components within the bilayerThe phospholipid molecules within the bilayer are in constant motion. Thepossible modes of mobility are displayed in Figure 4.6.Polar hecdgroup Fast lateralregion—flexing and diffusion inchanges in two dimensionsorientation of the bilayerRapid flexing ofocyl chainsKink formation inthe ocyl chains —o very rapidprocessFlip— flop —on extremely SlOw‘one-for-one’exchange processRapid rotational diffusionabout the long axis ofthe phospholipidFigure 4.6 Modes of mobility of the lipid components of the bilayer.80The freedom of rotation around the carbon-carbon bonds in a saturated fatty acidleads to an extended configuration that allows maximum intermolecular interactions tostabilize the structure. When cis double bonds are present, the chain becomes fixedpreventing efficient packing of the molecules, leading to less favorable interactionalenergy. The insertion of cholesterol into the bilayer restricts the motion within the bilayer(Figure 4.7).0—P—0—CH2 o — Motion MotionI I restricted unaffected-o o-cH2C-o0groupFigure 4.7 Insertion of cholesterol into a phospholipid bilayer. A schematicrepresentation of the alignment of a cholesterol molecule with aphospholipid in a lipid bilayer.1.1 nm O.SnmRigid sterol • Flezibtering tail814.1.5 Liposomes as models for biological membranesThe liposome was developed by Bangham et a!. as a model for biologicalmembranes.’27 They were the first to discover that the hydration of lipids results in theformation of sealed vesicles.128 In 1972, Cohen and Banghain made an extensive studyof the capacity of a large number ofnon-electrolytes to cross the membrane of suchvesicles.129 They found that the permeability of a molecule across the liposomalmembrane is highly dependent upon its size and its solubility in an organic solvent. Thus,they showed that the basic permeability properties of biological membranes are mimickedby these artificial lipid bilayers. In addition, because the composition of the bilayer can beprecisely controlled which allows liposomes to be formed with a similar membranecomposition to actual cells, studies have shown that liposomes are excellent models fortransport studies; they behave as usefhl models ofbiological membranes.’3°4.1.6 Types of liposomesThere are three types of liposomes: multilamellar vesicles (MLVs) which formspontaneously upon the hydration of lipid and contain multiple bilayers, small unilamellarvesicles (SUVs), which are less than 50 nm in diameter and contain a single bilayer andlarge unilamellar vesicles (LUVs), which are between 100 and 400 nm in diameter andalso contain only one bilayer.133 MLVs are unsuitable as models for membranes becauseof difficulties in reproducible production and variations in size. SUVs are also unsuitableas models for membranes because of the high degree of membrane curvature and minisculetrapped volumes. However, LUVs may be made reproducibly to specific sizes and arequite stable which makes them the ideal choice for a model membrane system.’3°824.1.7 Large Unilamellar Vesicles (LUVs)LUVs may be produced through solvent evaporation/vaporization,’34sonication,135 and extrusion techniques.126 The extrusion technique is the mostreproducible and was employed in this thesis. In the extrusion technique, the LUVs areproduced by passing MLVs through polycarbonate filters of defined pore size undermoderate pressure ( 500 psi). This is achieved by using a device known as, “TheExtrude?’ (Figure 4.8).126,136 The relatively high pressures allow homogeneously sizedLUVs to be produced ranging in size between 50 and 200 nm in diameter depending uponthe filter size.’37 Electron micrographs of vesicles produced after extrusion throughFigure 4.8 The Extruder.83different sized filters are displayed in Figure 4.9. The mean diameter and standarddeviation (S.D.) of the extruded LUVs is displayed in Table 4.1.137Figure 4.9 Electron micrographs ofvesicles prepared by extrusion through; A)400; B) 200; C) 100; D) 50; and E) 30 nm pore size polycarbonate filters.(from reference 137).84z1-4z4-4xwz4-4U)IzLi,Li,0Table 4.1 Mean diameter ofLUVs produced via the extrusion technique.Filter pore size (nm) Mean diameter±S.D. (nm) Mean diameter±S.D. (nm)Electron microscopy Light scattering400 243 ±91 not determined200 151±36 180±55100 103±20 139±3650 68±19 74±18The degree of unilamellarity of the extruded LUVs was tested by Mayer eta!.137They measured the intensity of the signal from the phosphorus molecules, on thephospholipids of LUVs produced by the extrusion technique, before and after the additiongo80706050400 2 4 6 8 IDNUMBER OF PASSESTest of the unilamellarity ofvesicles extruded the indicated number oftimes through 400 (.), 200 (.), and 100 (o) nm pore size filters.Figure 4.1085ofMn2by using31P-NMR. The Mn2broadens the spectroscopic signals of thephospholipids on the outside of the bilayer so that only the spectroscopic signals from thephospholipids on the inside of the bilayer remain. The results from this experiment are- displayed in Figure 4.10. If the phospholipid bilayer were completely unilamellar, only50% of the initial signal would remain after the addition of the Mn2.4.1.8 How is biomobility of environmentally sensitive compounds traditionallymodelled?In order to assess the environmental impact of a compound it is desirable to havesome measure of its ability to pass through biological membranes. When estimations ofthis type are needed octanol/water partition coefficients are often quoted.108 Forexample, the octanol/water partition coefficients for chlorobenzenes and chiorophenolscorrelates linearly with the uptake of chlorobenzenes and chlorophenols by trout, asshown by Neely el aL109 This correlation is generally useful because many of the organiccompounds which are either known to or suspected to constitute a hazard to theenvironment are highly lipid soluble° and, therefore, have relatively large olw partitioncoefficients (>102). These compounds usually exist as a single molecular species in theenvironment and do not possess ionizable groups. This is fortunate because a partitioncoefficient is defined as the distribution of a single molecular species between twophases. 1114.1.9 Difficulties in using partition coefficients to predict the biomobility of weakacidsWeak acids such as MM.A and DMA dissociate in solution producing two or threemolecular species. This makes partition coefficients difficult to determine because the86exact identity of the molecular species in each phase is rarely known. For example, in thepartitioning of an aliphatic carboxylic acid with a pKa of 4.5 and the aqueous phasebuffered at pH 8.5, only 1/10000th of the acid will be in the neutral form in the aqueousphase, and yet almost one-half of that present in the octanol phase will be the un-ionizedspecies.11’ Further difficulties arise because these hydrophilic compounds partition verypoorly into octanol and therefore, would have very small partition coefficients (<10-2).Thus, there are many difficulties in accurately determining the partition coefficients forweak acids. This uncertainty makes it quite difficult to correlate partition coefficients withthe bioaccumulation of weak acids.In general, an additional drawback to using partition coefficients to predictbiomobility and bioaccumulation for any molecule is that a partition coefficient is derivedfrom a system at equilibrium and, therefore, does not contain any kinetic informationwhich is necessary if the rate of uptake is to be predicted.4.1.10 The bioaccumulation of arsenicals by Candida humicolaThe uptake of arsenate by Candida humicola cells was investigated by Cullen etaL’ 12 At arsenate concentrations lower than 0.02 mM there seems to be a linearrelationship between the rate ofuptake by the cells and the external arsenateconcentration. If phosphate is added to the external solution at an equimolar amount tothe arsenate, the rate of arsenate uptake by the cells was reduced to about twenty percentof the rate in the absence of phosphate. If the cells were incubated with electron transportinhibitors, the rate of arsenate uptake by the C. humicola was decreased. Thus, it wasconcluded that the the uptake of arsenate by the cells required an active transportmechanism. Arsenate uptake has also been monitored for several other organisms, usually87with the objective of studying its effect on phosphate uptake113-124 and to establish itsrole as a metabolic precursor to more complex organoarsenicals.5The uptake of arsenite, MMA, and DMA by C. humicola cells was also measuredby Cullen eta!,’12 and there was no detectable uptake of these arsenicals within the first30 minutes of adding them to the C. humicola cells. Culturing the cells in the presence ofMtvlA and DMA did not affect their ability to uptake arsenite, MMA and DMA. Thus, itwas concluded that the uptake of arsenite, MMA, and DMA by the cells was via slowpassive diffusion.4.1.11 The bioaccumulation of DMA and MMA by unicellular organismsThere are considerable differences in the uptake of DMA and MMA by unicellularalgae. When grown for 10 days in a medium containing 2 ppm DMA Isochrisis galbanaaccumulated —76% of the available arsenic.’9 From media containing I ppm and 5 ppmMMA, the same cellular concentration ofI. galbana over the same time periodaccumulated —25% of the available arsenic. In a similar experiment, Dunaliella lertiolectaaccumulated —50% of DMA and —19% of MMA from media containing 0.5 ppm of theappropriate arsenical.Recent work by Hao Li’25 with the marine algae Acetabularia chftonhi has shownthat when grown in 10 ppm DMA for 7 days the algae accumulate 28.08 ppm DMA, whilethe same cellular concentration of algae over the same time period, grown in 10 ppmMMA, the algae accumulate only 6.48 ppm MMA. Hao Li observed similiar results fromthe same types of accumulation experiments with A. c4/ionii from a medium whichcontained 0.9 ppm of either DMA or MMA. In these experiments, algae accumulated20.39 ppm ofDMA and 1.56 ppm ofMMA.884.1.12 Scope of workAs discussed in Section 4.1.9 partition coefficients are a poor model to use topredict the bioaccumulation and biomobility of DMA and MMA, because of the inherentdifficulties associated with the measurement of partition coefficients for weak acids.Clearly, an alternate model is desirable and this new model would preferably contain akinetic component so that phrases such as “slow passive diffusion” which was used todescribe the entry ofDMA and MMA into the C. humicola 12 can be quantified.The diffusion coefficients across liposomal membranes should model the environmentalbiomobility and bioaccumulation ofweak acids more closely than partition coefficients Inthis chapter, this hypothesis will be tested by measuring both the diffusion coefficientsacross the liposomal membrane and the 0/w partition coefficients for MMA and DMA.894.2 THEORYThe following equations are used to describe the one directional efflux of[3H]labeled MMA and DMA from LUVs which were labeled with[14C]. The permeant wastrapped inside the LUVs, a concentration gradient was established so that the permeantdiffused from the inside to the outside of the LUVs, and the amount of radiolabeledpermeant remaining inside of the LUVs was measured at specific time intervals.4.2.1 The rate constant, permeability coefficient and activation energyThe results in this chapter were obtained by means of efflux measurements wherethe rate of diffusion out of the LUVs in response to the concentration gradient ismonitored. Efflux studies require less radioactivity and it has been established that theresults obtained are essentially the same as from influx (where the rate of diffusion into theLUVs is monitored) measurements.’38 Assuming that the permeation of arsenicals acrossthe liposomal membrane follows first order kinetics and that the influx of molecules isnegligable, the following equation can be written for efflux;-ÔXJÔt=ic(X) [4.1]where X is the concentrations (molsfL) of the radiolabeled compound inside the LUVs attime t and is the first order rate constant (s1). Integration gives;ln((X0)/(X)) = ict [4.2]where X€ is the concentration of the radiolabeled compound inside the LUVs at t =0-lnX0+lnX=-kt [4.3]X and X0 are concentrations and are proportional to the amount of permeant trapped inthe LUVs where, the permeant and the liposome concentrations are represented by the 3Hand the 14C labels, respectively; thus90X = cx(3HJ’4C)t and X0 = cx(3H114C)0 [4.4]It is now possible to write:-in a(3H/14C)0+ in cx(3H1’4C)t = -kt [4.51which leads to;ln(3HJ’4C)t=-kt+C [4.6]A plot of in(3HJ14C)t versus t will yield a slope equal to -k and an intercept equalto(3HJ14C)0.From the first-order rate constant Ic; the permeability coefficient P (cm s-i) can becalculated if the area and the trapped volume of the LUVs are known. 138P = k x Volume/Area [4.7]Assuming that the average size of an EPC head group is 60 Angstroms2and that asingle biiayer is formed, the area is calculated to be 1.81 xl cm2 (pmoi lipid)-’. Finallythe activation energy for the diffusion can be calculated with the aid of an Arrhenius plotaccording to equation [4.81:P = A exp(-EaIRT) [4.8]where; P = permeability, A = constant, Ea = activation energy and R and T have theirusual meanings.914.3 EXPERIMENTAL4.3.1 Scintillation CountingThe[3H]- and[14C]- activity was measured by using a Packard 2000 CA liquidscintillation counter. Packard software was used for a simultaneous determination of bothlabels. Typically, 100 i.1 of sample was added to a 20 ml glass scintillation vial whichcontained 800 p.1 of deionized water and 9 ml of scintillation cocktail. The samples werecounted for two minutes.4.3.2 Chemicals and ReagentsEgg phosphatidyl choline (EPC) and[14C]-dipalmitoylphosphatidyl choline(DPPC) were purchased from Avanti Polar Lipids, Birmingham, AL, USA, and Du PontCanada, respectively. The synthesis of the[3Hj-MMA and[3111-DMA used in thischapter is described in chapter 6.4.3.3 Preparation of LUVs and Sampling Technique.Dry EPC (75 mg) with[14C]-DPPC as the label was hydrated with buffer [1 cm3,20 mmol dm3 Hepes(C8H18N204S)and 150 mmol dm3 NaCI adjusted to pH 7.41containing either 17 mg of[3Hj-MMA or[3Hj-DMA. The resulting multila.mellarliposomes were subjected to five freeze-thaw cycles employing liquid nitrogen to enhancesolute distribution.’39 The solution was then transferred to a device, Figure 4.8, (SciemaTechnical Services Ltd, Richmond, B.C., Canada) that was used to extrude multilamellarliposomes through 100 nm pore size filters (Nucleopore Inc.) under 300 psi (2000 kPa).After ten such extrusions LUVs were produced.’26 The untrapped compound wasremoved from the LUVs by passing the solution through a Sephadex G-50 column (1.592cm x 15 cm) that had been pre-equilibrated with the buffer. The LUVs (1.5 ml) werecollected, diluted to 4.5 ml, divided into three equal portions and placed in a constanttemperature bath. This point in the experiment was designated as time zero. Thesampling proceedure is summarized in Figure 4.11. At appropriate intervals of time 100 i1 was withdrawn and loaded onto a dry Sephadex G-50 column packed into a 1 cm3disposable syringe, and the LUVs were eluted by centrifugation at 2000 rpm for 2minutes. Deionized water (800 .il) was added to the eluant along with Aqueous CountingScintillant (ACS) (9 ml) from Amersham and the3HJ14Cratio was determined byscintillation counting.1) withdraw sample4)repN%2) add sample to spin columncount eluted LsFigure 4.11 Radiolabeled sampling procedure.934.3.4 Determination of Octanol/”Vater Partition CoefficientsEither[3H]-MMA or[3H]-DMA (4 mg) was added to buffer pH 7.4 (50 ml) andoctanol (50 ml) in a volumetric flask. The flask was stoppered and submersed up to itsneck in a thermostated bath set at 25 °C. Each flask was vigorously shaken every 5minutes. After 30 minutes the two phases were separated by using a separatory funnel and1 ml from each phase was withdrawn to be counted. The ratio of the counts was used todetermine the partition coefficient.944.4 RESULTS AND DISCUSSION4.4.1 Rate constants and permeabiities for the diffusion of MMA and DMA acrossliposomal membranes (100 nm LUVs at pH 7.4)Buffered solutions ofMMA and DMA (pH 7.4) were used to hydrate dried lipidwhich was then extruded through 100 nm sized filters to form LUVs. A concentrationgradient was established and the efflux ofboth molecules across the liposomal membranewas measured and plotted according to equation 4.6. The rate constants were determinedfrom a linear regression of these plots which are shown in Figure 4.12.The permeability coefficients were calculated from the rate constants and thephosphorus concentrations according to equation 4.7. The rate constants and permeabilitycoefficients for MMA and DMA at three different temperatures are displayed in Table 4.2.Table 4.2 Rate constants and permeability coefficients for MMA and DMA at pH 7.4across the liposomal membrane (100 nm filter size).Compound Temperature (°C) Rate constant (s1) Permeability (cmls)MMA 22 1.4 (± 0.2) x i0 1.2 (± 0.3) x 10-13MMA 31 6.3 (± 0.9) x i0 5.5 (± 1.1) x iO-13MMA 39 2.1 (± 0.3) x 10-6 1.8 (± 0.4) x 10-12DMA 24 4.3(±1.1)x105 3.7(±1.1)x10DMA 32 2.3 (± 0.6) x 10 2.0 (± 0.6) x i010DMA 36 2.8 (± 0.7) x 10 2.5 (± 0.8) x 10-1095-0.5L)r-)—1MMA RATE CONSTANT DETERMINATIONTime (hours)DMA RATE CONSTANT DETERMINATION35090Figure 4.12 Efflux measurements ofMMA and DMA. Data are plotted according to22°C0—0.924°C32°CTime (minutes)equation 4.6.96The results which are displayed in Table 4.2 show that the rate constants and thepermeability coefficients for DMA are much greater than for MMA. At 25°C theinterpolated values of the permeability coefficients across the liposomal membrane forDMA and MMA are 4.5 x 10-11 cm r1 and 1.4 x i013 cm s, respectively. The onlystructural difference between the permeants is the substitution of a -CH3 group for an-OH group. The data support the notion that a hydroxyl group added to a molecule willdecrease the permeability 100-1000 fold, while adding a methyl group will increase thepermeability 5 fold.14° Also, MMA has pK1 = 4.58 and pK = 7.82, whereas DMA haspK =6. 19. At a pH of 7.4, which is the pH of the uptake studies described in Section4.1.3, —95% ofthe DMA is present as an anion with a single negative charge; theremaining 5% is neutral. At pH 7.4 approximately 75% of the MMA is present as ananion with a single negative charge and the remaining 25% is doubly negatively charged.4.4.2 Activation energy for the diffusion MMA and DMA across liposomalmembranes (100 nm LUVs at pH 7.4)The natural logarithms of the permeability coefficients were plotted versus thereciprocal temperature (Figure 4.13). The activation energies for the diffusion of MMAand DMA across the liposomal membranes were determined from a linear regressions ofthe plots according to equation 4.8. The results are displayed in Table 4.3.Table 4.3 The activation energy required for diffusion of MMA and DMA throughthe liposomal membrane at pH 7.4.Compound Activation Energy (KJ mo11)MMA 220±35DMA 130±3097—I4MA ACTIVATION ENERGY DETERMINATON0.0034DMA ACTIVATION ENERGY DETERMINATIONlIT0.0034Figure 4.13 Activation energy determinations for MMA and DMA. Data are plottedaccording to equation 4.8.ClIT00.003298The activation energy for the diffusion of a molecule through a membrane is theenergy required for the molecule to enter into the phospholipid bilayer from the aqueousphase. The differences in the activation energy for MMA and DMA could partially beexplained by the fact that MMA has a greater hydrogen bonding capability with thesurrounding water molecules than does DMA. Indeed, it has been shown in previousstudies with erythritol, glycerol and glycol, that the activation energy for permeationthrough artificial membranes corresponds directly with the number of hydrogen bonds thatneed to be broken in order for the permeant to undergo complete dehydration. 1414.4.3 Octanol/Water partition coefficientsEither[3H]-MMA or[3H]-DMA was added to buffer and octanol and the ratio ofcounts in the two phases was determined giving the octanol/water partition coefficients forboth compounds. The results are displayed in Table 4.4.Table 4.4 The octanollwater partition coefficients for MMA and DMA.Compound Octanol/water partition coefficientsMMA 7.4(±1.5)x103DMA 8.4 (± 1.7) xThe two partition coefficients are essentially the same, whereas the permeabilitycoefficients vary by a factor of 330. To a first approximation permeability coefficients andpartition coefficients are related by Overton’s rule which states that one is proportional tothe other.’42 However, in practice the situation is more complex as factors such aspermeant size, shape, charge and hydrogen bonding capability, all affect the relationship.99Charged species do not significantly partition into organic solvents which is why thepartition coefficients for both molecules are quite small. For comparison purposes glucosehas permeability and octanol/water partition coefficients of 3.0 x 10-11 cm s’ and 1.0 xrespectively. The permeability’38ofwater is 4.4 x i0 cm s with an activationenergy of 33-38 kJ moP’ and an octanol/water partition coefficient’43of 0.041.4.4.4 SummaryThe permeability coefficients through the liposomal membrane (1.4 x 10-13 cmand 4.5 x 10-11 cm s’ for MMA and DMA, respectively) provides a better indication ofthe environmental mobility for DMA and MMA than do octanol/water partitioncoefficients (7.4 x i0 and 8.4 x i0 for MMA and DMA, respectively). Based uponpartition coefficients one would expect a similiar biomobility for both DMA and MMA.This is clearly not the case, as shown by the results in Section 3.3.2, (where DMA is takenup by microscopic organisms and biotransformed much more rapidly into arsenobetainethan is MMA) if the rate determining step is assumed to be the entry of the either DMA orMMA into the microscopic organisms. The results of the permeability coefficient wouldlead to the prediction that a cell would be more likely to accumulate DMA than MMA, ifthe only mechanism of entry into the cell is via passive diffusion (Section 4.1.4). Thissupposition is in agreement with the results described above for I.galbana andD.tertiolecta (Section 4.1.5). In addition, the use of liposomes has enabled aquantification of the phrase “slow passive diffi.ision”. The permeability coefficients alsocontain a kinetic component which gives an indication of the rate ofuptake which islacking from partition coefficients. These ideas are fi.irther investigated in the nextchapter.100CHAPTER 5THE DEVELOPMENT OF AN NMR TECHNIQUE TO MEASURE TilE RATEOF DIFFUSION THROUGH LUVs AND ITS APPLICATION TO MMA ANDDMA5.1 INTRODUCTION5.1.1 Traditional techniques used to measure the diffusion coefficients across themembranes of LUVsThe permeability coefficients for various molecules (neutral and charged) acrosslipid bilayers such as those found in LUVs and other types of membranes can be measuredby using several different techniques.146 These techniques require measurements of thechange of concentration of the permeant on either side of the membrane as a function oftime. The sampling proceedure typically involves withdrawing small aliquots from thebulk solution of LUVs (as in Chapter 4) and separating the permeant molecules on theoutside of the LUVs from those on the inside of the LUVs. The separation can beachieved by using chromatography,154millipore filters,155 extraction and variouswashing techniques. Usually the permeant is labeled with radioactivity or is fluorescent sothat the concentration after separation can be measured.LUVs and cells also shrink or swell in response to changes in the osmotic pressureacross the membrane.’46 This change ofvolume can be measured by using lightscattering techniques which allows the rate of the flow ofwater across the membrane tobe measured.’56 Also, since the rate of flow of permeants across the membrane alters theosmotic pressure, the rate of permeation of substances other than water can bedetermined.1015.1.2 Spectroscopic techniques used to measure the diffusion coefficients acrossthe membranes of LUVsSpectroscopy may be used to measure the diffi.ision of molecules acrossmembranes if the spectroscopic signals of the permeant can be differentiated on either sideof the membrane. In NMR spectroscopy, this can be accomplished through the use ofshift or broadening agents.157 The area of a spectroscopic signal is directly proportionalto the number of molecules which generate the signal. As the permeant molecules diffuseacross the membrane, the corresponding spectroscopic signals on either side of themembrane will change in intensity and the rate constant for the diffusion process can bedetermined by measuring the rate at which these spectroscopic signals change. The rateconstant can then be correlated to the permeability coefficient for the difflising molecule.For example, the permeability coefficient for Na+ crossing the membrane ofLUVswas determined by using23Na-NMR spectroscopy158 and the shift agent Dy(PPPi)27.In addition, Prestegard et al used NMR spectroscopy in conjunction with Pr+3 tomeasure the permeability coefficients for maleic acid159 crossing the membranes of LUVsproduced by sonication.The osmotic permeability has also been measured by using the NMR transverserelaxation times (T2) of the extra-vesicular water and the intra-vesicular 60 Ifwas added to the outside of the LUVs the transverses relaxation times of the wateron the inside and the outside would be different. Cafisio and Hubbel have used ESRspectroscopy to follow the transmembrane movement of the spin-labeled phosphoniumion.’6’ Thus, the use of using magnetic resonance spectroscopic techniques to measurethe diffusion coefficients across membranes is not a new one, however, work in the field islimited to a few select molecules.1025.1.3 Advantages of using spectroscopy over traditional techniques to measure thediffusion of molecules across membranesSpectroscopic techniques, such as NMR and ESR permit the diffusion ofmolecules across membranes to be monitored without a physical separation thuseliminating the sampling procedure. Instead of the labour intensive procedure used inChapter 4 (see Figure 4.11) the sample simply remains in the magnet and spectra arerecorded and stored automatically. In addition, the system is closed and nothing is addedto or withdrawn from the media during the diffusion of the permeant. This is unlike theprocedure in Chapter 4 where samples were continuously withdrawn from the bulk media.Also, because the system is closed, the experiments can be monitored until equilibrium isreached enabling the trapped volume to be determined (see Section 5.3.7). The absence ofradioactivity decreases the costs of the permeant and eliminates the scintillation vials andfluid and improves the safety of the working environment. Also, the elimination of thedisposable spin columns which were composed of a syringe and some Sephadex (as usedin Chapter 4) reduces the costs substantially.5.1.4 Difficulties in using NMR spectroscopy to measure the diffusion of moleculesacross membranesThe most difficult part of using NMR spectroscopy to measure the diffusion ofmolecules across membranes is in finding a suitable shift agent. The shift agent must beable to shift the spectroscopic signal while not reacting with the permeant molecules orwith any other molecules in the system. The shift agent must also be impermeable to themembrane and, in addition the spectroscopic signals from the permeant should notresonate at the same position as any of the other molecules in the system. The bestpermeant molecules for this technique have fairly simple NMR spectra with a singlet that103can be shifted, because it is difficult to shift doublets and triplets so that they are properlyresolved. The other important limiting factor is the rate at which the permeant moleculesdiffuse across the membrane. If the permeant diffuses with a permeability coefficientgreater than 10-8 cm s’, then it becomes difficult to measure the coefficients accuratelywithout using T2 measurements. There are also solubiity limitations on the permeant.There must be enough permeant in solution so that the spectroscopic signal can be clearlydetected.5.1.5 Scope of the workIn this chapter, a NMR spectroscopic technique was developed to measure thediffusion coefficients for molecules across liposomal bilayers. The NMR spectroscopictechnique employed1H-NMR with Mn2 as a shift and broadening agent, LUVsproduced by the extrusion technique and both MMA and DMA as the permeantmolecules. The development of this technique allowed further investigation of thepermeation of MMA and DMA across the liposomal membrane. The effects of changes tothe pH, temperature, liposome size and membrane composition on the rate constants werestudied. In addition, the theory to properly describe the diffusion across the sphericalliposomal membranes was developed and equations to describe the processes werederived.1045.2 THEORY5.2.1 What causes molecules to diffuse across membranes?There will be a net diffusion (transport) of molecules across a membrane if there isa chemical potential difference across the membrane.ie =Outrntj [5.1]where i.t is the chemical potential and is defined as;= *+RThi++V [5.2]where a is the activity, c is the membrane potential and p is the osmotic pressure.If the osmotic pressure and membrane potential are equivalent on both sides of themembrane and there is a difference in chemical potential (A 0), then this difference iscaused by an unequal activity across the membrane. Activity is related to concentration bythe following formula;a=y±C [5.3]where is the activity coefficient and C (mol m11)is the concentration.The net flux across a membrane is defined as;’46JA = DAÔC/ôx DAC/x = DA(C0h1tCm)/Ax [5.4]where 3 (mol cm2 s_1) is the flux, A (cm2)is the surface area, D (cm2s4) is thediffi.isivity coefficient and x (cm) is the membrane thickness.105A schematic diagram of the net flux across a membrane in response to a chemicalpotential difference is displayed in Figure 5.1.IOutside Inside(Cj < Cout)FluxFigure 5.1 Flux across a membrane in response to a difference in chemical potential.1065.2.2 Diffusion from a spherical vesicle.The net flux of molecules across a sphere may be described by Ficks LSt Law as:JA = DAaC/ar = D4itr2ôC/ô= ôn0t/ôt [5.5]where A (cm2)is the area, r (cm) is the radius, D is the difihisivity coefficient (cm2s1)and Vj (ml) is the trapped volume of the sphere, while C (mol mi-I) is the concentration,and n is the number of moles or molecules.The net flux may also be written as;JA= ôflOut/& = -âniniôt = kt(nmjnfVmin - nmout/Vffout)= klK(naqjfVaq- naq0/V)= k(flaqNaq - naq0/V [5.6]where and nmout are the number of molecules or moles of permeant on the surfaceof the inside and outside of the sphere, respectively, naq1 and na%ut are the number ofmolecules in solution on the inside and outside of the sphere, respectively, K is thepartition coefficient (K = cm/caq), k’ (cm3 s-i) is the rate constant within the membrane,k (cm3s1) is the rate constant across the membrane and equals k’K and V is the volume(cm3). The equation is written in terms of number of molecules or moles instead of themore traditional concentrations because NMR spectroscopy provides a direct measure ofthe number of particles rather than a concentration measurement. The above terms aredisplayed schematically in Figure 5.2.vaq0, 107Figure 5.2 Schematic depiction Of Vaq, Vaq0Vmin, Vm0D, K, k’, k,[J&1m0t [J{4]m pjaq0 [I{]aqFicks 2nd Law states that:ãC/ãt = (D1r)ã/ôr(rôC/ãr)At equilibrium (ÔC/ôt = 0):ô/ãr(r2ãC/ãr) =0r2ôC/ãr = ewhere e is a constant, therefore,8C/ãr = e/r2[5.7][5.8][5.9]I[5.10]108[5.11][5.12][5.13][5.14][5.15]Integrating 5.10 gives;C = -elr+bcmm = -e/rj+bCmout = e/rout+bcm- Cm0= -e(l/rm - i/rout) = -a((rj- rout)/rjnrout)e = - Cmout]rjnrout![rjn- rout])m r’mSchematic depiction of C‘- out’ rin and rout.Figure 5.3Substituting equation [5.10] into equation [5.5] gives:JA = ADÔC/ôr = 4nr2De/r = 4itDeSubstituting equation [5.15] into equation [5.16] gives:JA =47cDrjnrout[CmjnCmout]/(rinrout)[5.16][5.17]109JA=47tDrjnrout[nminNjn- nmoutfVout]/(rm- rout) [5.18]From equation [5.6]JA = k[nmmfVm -nm0tN1Therefore, rearranging equations [5.6] and [5.18] to solve fork’ gives:k’ =4lrrinroutD/(rin-rout) = AD/Ax [5.19]Thus, the rate constant is dependent upon the diffusivity coefficient within themembrane and upon the area and the thickness of the sphere. Large Unilamellar Vesicles(LUVs) are spherical.5.2.3 The Permeability Coefficient and Activation EnergyThe permeability coefficient contains three terms that are very difficult to measurein biological membranes: Ax the thickness of the membrane; D the diffusivity coefficientwithin the membrane and K the partition coefficient of the permeant between themembrane and the aqueous solution. The combination of these three terms given inequation 5.20, is defined as the permeability coefficient (P in units of cm s-i). P isrelatively easy to measure.PDK/Ax [5.20]Rearranging equations [5.19] and [5.21] and solving for P gives:P=WKJA=k/A [5.21]The activation energy for diffusion may be calculated by using equation [4.8] from Section4.2.1 of the previous chapter.1105.2.4 Derivation of the Equations used to describe the number of particles oneither side of the membraneThe equations used to describe the number ofparticles on both the inside and theouside of the LUVs at any time during the experiment are derived after the followingassumptions are made. The rate of diffusion into the LUVs (influx) will equal the rate ofdiffusion out of the LUVs (efflux). The LUVs are of uniform size and the volume doesnot change during the course of the experiment (i.e., the conditions on both sides of themembrane are isotonic). The activity coefficients have values of I and, therefore, theactivities of the acids are equal to the concentrations. The dissociation of the acid is rapidwith respect to the time of diffusion so that the acid dissociation can always be consideredan equilibrium process. Assuming that the permeant does not decompose or react withany of the molecules which are present.The net flux of molecules may be written as:&hin/ãt = ouil8t k(ninNinnoutNout) [5.22a]ôCfôt= k(CjnC0ut) [5.22b]where n is either the number of mols or the number of particles and C is the concentrationof the permeant in the aqueous solution on either the inside or the outside of themembrane, respectively. k (ml s1) and ç (r’) are both rate constants to describe thediffusion of the permeant.Let:N = nojfl+noout = neq+neq0= ntin+ntout [5.23]where N is the total number ofmolecules and n0, eq and are the number of moleculesinitially, at equilibrium and at any time during the experiment. N is a constant because thenumber of molecules in the NMR tube does not change.111Also let:V’ = (l/Vjn+lNot) [5.24]Substituting equation [5.23] into equation [5.22a] gives:outt = k((Nflout)Ninnout/Vout) [5.25]= k(NNinnout(lNin+1/Vout)) [5.26]Substituting equation [5.24] into equation [5.26] gives:61)out’t = k(N/V - noutV’) [5.27]Rearranging and integrating equation [5.27] gives;outW’b”in - noutV’) = kãt [5.28]- 11out’7’) fltno = V’kt [5.291There will be some permeant molecules outside of the LUVs at time zero in theexperiments (ie. n°out 0) therefore:1fl(VinfltoutVI)1(NNinflOoutV)) = V’kt [5.30]ln((NntoutVIVin)/(NnOoutVVin)) = V1kt [5.31]Let:f VjnN [5.32]where f is the ratio of the volumes inside and outside the LUVs.112Substituting equation [5.32] into equation [5.311 gives:ln( 1 ...(nt rI1t tf)1N)4n( 101°out-11°outf)IN) = -V’kt [5.33]Let the number of particles inside and outside of the LUVs at equilibrium be written as:neq0= N/(1+f) [5.34]and neq=N/(1+(1/f)) [5.35]Substituting equation [5.34] into equation [5.33] gives:ln( 1 +f))-ln( 1 1 +f)) = -V1kt [5.36]ln( 1 ntout/ne%ut)1n( 1 = -(1+f)kt/Vj [5.37]= -(1 +f)ktfVj [5.38]Equation [5.38] may be rearranged to yield equation [5.39] which gives the number ofparticles outside the LUVs at any time t:ntout = [5.39a]or = [5.39b]where k= k/VjSubstituting equations [5.23] and [5.35] into equation [5.38] and rearranging givesequation [5.40] which gives the number of particles inside the LUVs at any time t:ntin = neq+(nOneq)e-(l+f)ktN [5 .40a]or nt1 = [5.40b]1135.2.5 Permeation of a Weak AcidFor a molecule such as DMA, which is comprised of two molecular species insolution (as shown in Figure 5.4), the overall flux ofmolecules from the LUVs will be acombination of the two species. This combination may be written as;0 0II_____________IIH3C—As—OH+H20 H3C—As—O-+H3OI ICH3 pK = 6.28 CH3Figure 5.4 The two chemical species ofDMA in aqueous solution.afljn/at=ãflJjj/ôt + Ain [5.41a]or ôfl/ôt = + kA..(flA..inNjflA..out/Vout) [5.41b1where the subscripts AM and A- refer to the neutral and anionic forms of the weak acid.Let:n&j = cn and A-= (1-cz)n [5.42]114where n is the total number of particles and cx is the degree of dissociation of the weakacid and equals KaJ(Ka+[H+]) for a monoprotic weak acid andKaJ([Hj2+K1[Hj)for a diprotic weak acid.Substituting equation [5.42] into equation [5.41] gives:-ãn/ãt = + kA((l)njNj(1cX)floutNout) [5.43]-ãn/ôt = ctkpji(njnNin-nout/Vout) + (1cx)kA..(ninNinnout/Vout) [5.44]If cxk&j.j is equal to (1-cx)kA the rate of overall flux of molecules will beindependent of pH, however, if cxkp.j is greater than (1-a)kA the rate of overall flux ofmolecules will be pH dependent.The methyl resonance of DMA, a weak acid, is composed of both the neutral andthe anionic molecules. The position of the combined methyl resonance for the neutral andthe anionic forms of DMA varies slightly depending upon the percentage of each chemicalspecies. The integral of the combined methyl resonance is related to the number ofparticles of each chemical species by constant scaling factors such that:‘total =1DMAHDMAH + IDflDK [5.45]where ‘total is the total integral and1DMAH ‘DMA- are the scaling factors.The rate of change of the total integral may be written as:=1DMAHDMAH’ + IDMADMAJ& [5.46]If the anionic form of the molecule does not significantly permeate the membrane thenk&J{ >>>kA.... For this sitation, ôflA&J/ãt>>> A-in’’ and (1CL)kA..(flinNin1ouilVout) (equation [5.44]) will be approximately equal toO. Thus,(equation [5.46]) will also be equal to 0. Therefore, the equations used to describe the115rate of change of the total integral may be written as;rout = Ieq0+(IO4eq01)e(l+f)CLkA}{t/Vjn [5.47a]or rout = Jeq0+ [5.4Th]and it = jeq + [5.48a]or it = ieq + (IOIeqj)e(l+f)c4cAHt [5.48b1Thus, the rate constant will vary linearily with the [Hj since:[DMAH] = [DMA1[Hj/KA [5.49]Thus,k=ckpj1 [5.50a]icc4cM{ [5.50b]and P=cxPpj.1 [5.50c]where k, and P are the pH dependent diffusion coefficients; while icM{, and Pjare the diffusion coefficients for the neutral species of the weak acid.5.2.6 Comment on D, P, k’, k,, kAB, hAH and AI{To avoid confusion later the diffisivity coefficient (D in cm2r1) and thepermeability coefficient (P in cm are properties of the membrane and the permeantand are independent of liposomal size, trapped volume and surface area. They are relatedby the following formula;P=DK/Ax [5.52]where Ax is the thickness of the membrane and K is the partition coefficient.116The rate constant for the permeant within the membrane (k’ in cm3 s-i) is definedin Section 5.2.2 as:kDA/Ax [5.53]The rate constant across the membrane (k in cm3 s4) is related to the rateconstant within the membrane by the partition coefficient such that:k = kK = DAKJAx = PA [5.54]Since the trapped volume is incorporated into k, it will scale only with the surface area ofthe LUVs. If unilamellarity is assumed, and the same amount of lipid is present in eachexperiment, k will be independent of the size of the LUVs.The traditional first order rate constant (k in s’) is dependent upon the size of theLUVs and is proportional to hr. It is defined as:k=k/V=PAIV=DAKNAx=3DK/rAx [5.55]where r is the radius of the LUVs.Weak acids or bases are present in aqueous solution as neutral and as chargedmolecules whose ratio will vary with pH. The pH apparent rate constant will be acombination of the rate constants for both the neutral and charged molecules. The rateconstant for the neutral form of the molecules is designated as kj (cm3 s) or ic&jj(s-i) and is independent of the pH. The permeability coefficient follows the same patternand Pj.j is the permeability coefficient for the neutral form of the molecule.1175.3 EXPERIMENTAL5.3.1 InstrumentationA Bruker AM 400 NMR spectrometer was used to acquire the spectra. Thesamples (0.6 ml) were placed in high precision NMR sample tubes (178 mm x 5 mm(outer diameter)) from Norell, Inc., Landisville, New Jersey USA. The water signal wassuppressed by using a presaturation pulse as shown in Figure 5.5. The first pulse is at thefrequency of the water signal which becomes saturated. The second pulse is over a muchwider range of frequencies and is implemented while the water signal is still saturated.Saturation pulse selective to H20timeAcquisition pulse-0’Figure 5.5 The pulse sequence used for water suppression.118The program which operates the spectrometer in the water suppression and thevarious delay modes is displayed in Figure 5.6. This program was designed toautomatically aquire spectra at specific time intervals during the diffusion experiments.JOHN FILE:JOHNJOHN:AU PROGRAM FOR DIFFUSION EXPERIMENTSII2 ZEVD1D1HGGO=1 DOWR #1IF#1IN=2EXITVD VDLIST.OO11-25 = APPROPRIATE DELAYSFigure 5.6 The microprogram designed to operate the spectrometer.The delays between sample acquisition were varied depending upon temperature,pH, membrane composition, and permeant molecule. To establish the best set of delays it119is necessaiy to have some idea about to the length of the experiment until equilibrium isreached. The delay may be calculated according to the following formula:VD(1 )+(D1+AQ)+(NS+DS)+..30ms = delay [5.56]where VD is the variable delay, Dl is the duration of the presaturation pulse (0.5seconds), AQ is the acquisition time (1.016 seconds), NS is the number of scans (100),and DS is the number of dummy scans (2). The easiest way to determine the delay was totime the spectrometer with a stopwatch as it pulsed the required number of times.The spectra were acquired by using a pulse width of 6 milliseconds. A linebroadening term of 10 Hz was used during the Fourier Transformation. The resonanceposition of TSP which was added to the external solution was used to reference thespectra to 0 ppm.5.3.2 Chemicals and ReagentsEgg phosphatidyl choline (EPC) was purchased from Avanti Polar Lipids, inBirmingham, Alabama, USA, while the sodium salts6 1120 of both DMA and MMA werepurchased from the Fisher Scientific Company, Fair Lawn, New Jersey, U.S.A.5.3.3 Determination of the pKA’S for DMA and MMAThe pK&s for both DMA and MMA were determined by using the followingprocedure. Either MMA or DMA (3.5 g) was dissolved in deionized water (100 ml). Analiquot of this solution (25 ml) was withdrawn and placed into an Erlenmeyer flask (250ml) which was stirred. HC1 (0.1 M) was slowly added to this solution and the resultingpH was monitored by using an Orion pH Meter with a Fisher electrode which had beencalibrated with standard sloutions at pHs 4, 7, and 11. These titrations were performed intriplicate.120The1H-NMR resonance position of the methyl group on both DMA and MMAwas also measured as the pH was varied from 2 to 11 in increments of 1 and the data wastabulated and plotted.5.3.4 Preparation of LUVs and Sampling TechniqueThe following experimental conditions were employed for the diffusionexperiments that used N14R spectroscopy to measure the rate of efflux of both DMA andMMA across the liposomal membrane. DMA or MMA (50 mg) was dissolved in 2 mlHepes (300 mmolar in D20) and the pH of the resultant solution was adjusted by usingeither NaOH or HC1. This solution was added to dry egg phosphatidyl-choline (0.4 g) andvortexed to produce a milky white solution. The egg phosphatidyl-choline (1 g) wasdissolved in chloroform (10 ml) and stored in a freezer. The chloroform solution (2 ml)was withdrawn and pipetted into a glass test-tube and most of the chloroform wasremoved by blowing a gentle stream of nitrogen gas over top of the solution. Theremaining chloroform was removed by placing the test-tube in a vacuum apparatus for atleast 4 hours. The milky white solution was transferred to a plastic test-tube that wassealed with a screw on cap. This test-tube was submerged in liquid nitrogen until frozen(10 s) and was then submerged in water bath (50°C) until it had thawed (4 minutes). Thisprocess (freeze-thawing) was repeated 5 times to increase the unilamellarity of thephospholipid bilayer. The freeze-thawed lipid was added to an extrusion device (Figure4.8) which contained 2 polycarbonate filters. The filters had pore diameters ranging from50 to 200 nm depending upon the size of LUVs which were being used in the experiment.A nitrogen tank was attached to the extrusion device and the lipid was forced through thefilters with a pressure of between 200 to 500 psi. The extrusion proceedure was repeated10 times to ensure that all of the Large Unilamellar Vesicles (LUVs) which were producedby this technique were approximately the same size. The 2 ml ofLUVs were divided into2 equal aliquots so that the experiments could be performed in duplicate. The LUVs (1121ml) were added to a Sephadex G50 column (1.5 cm x 4 cm) and were eluted by usingHepes buffer (40 mmolar in H20 at the appropriate pH). This proceedure removes mostof the DMA that was not encapsulated and establishes the desired concentration gradientfor diffusion studies. Approximately I ml of the diluted vesicles were collected and atimer was started, as this was designated as time 0 in the experiments. The eluted LUVs(400 p1) were added to a NMR tube which already contained Mn2 (40 p.1 of 30 mmolarinD20), Hepes (135 p.1 of 40 mmolar inD20), TSP (25 p.1 of40 mmolar in D20) andeither glucose (28 mg) or NaC1 (4.6 mg). The water supressed1H-NMR spectra (25)were obtained at the appropriate time intervals by using a Bmker AM 400 spectrometerwhich had been programmed with the micro-program listed in Figure 5.6.5.3.5 Concepts to consider when designing these experimentsThese experiments require careful setting up because there are a number ofparameters which can be varied to produce optimum results. The initial permeantconcentration and buffer concentration inside of the LUVs must be set up so that the pHinside of the vesicles after the permeant has diffused out is relatively small (0.3 units ofpH). If the buffer concentration is too high with respect to the penneant concentration thespectroscopic signal from the buffer will overwhelm the signal from the permeant. Areally high buffer concentration also causes the spectrometer to select a low receiver gainwhich decreases the sensitivity as well. One way to avoid some of these difficulties is tomake the buffer concentration on the outside of the LUVs about 10% of the concentrationof the inside. The buffering capacity on both sides will still be the same because of thevolume difference. The buffer should also be relatively impermeable to the membraneover the time scale of the experiments.It is important to have a large trapped volume (10% ofthe total volume) so that122the initial permeant concentration produces the maximum signal on the spectrometer andthat the equilibrium position is easily measurable. To ensure a large trapped volume themaximum amount of lipid (O.2g per ml) which is easily extruded is generally used and thelength of the Sephadex column is kept fairly short (1 .5x5cm), to avoid a significantdilution of the LUVs.The osmotic pressure on both the inside and outside of the LUVs should be fairlyconstant so that the LUVs do not burst or shrivel up. To prevent this from happening arelatively impermeable substance (compared to the permeant) needs to be added to theoutside of the LUVs.The spectroscopic shift of the permeant is dependent upon the ratio of theconcentration of the shift agent to the permeant and upon the pH of the surroundingsolution. Finding the appropriate shift agent requires patience as different shift agentstend to shift different types ofcompounds.147 Success cannot be guaranteed.3-(Trimethylsilyl) proprionic-2,2,3,3-d4acid, sodium salt (TSP) which is oftenused to reference the spectra sometimes reacts with the buffer or the permeant and shouldbe eliminated or replaced in this situation with another spectral reference material. Thesignal to noise ratio increases with number of scans and, therefore, the number of scansshoud be maximized depending upon the time the permeant takes to reach equilibrium.5.3.6 Bligh-Dyer ExtractionThe phospholipid concentration of the post-column LUVs was determined so thatthe permeability coefficient could be calculated (see Section 5.3.9). Because ofinterference the sample must be free from arsenic when UV-visible spectrometry is usedfor phosphorus determinations. Therefore, the arsenic and the phospholipids must be123separated and this separation is achieved by using the Bligh-Dyer extraction proceedure.In this proceedure, the LUVs (0.5 ml) were diluted to 1 ml by adding deionized water.Methanol (2.2 ml) and chloroform (1 ml) were added to the diluted vesicles and thesolution was vortexed to produce a single phase mixture. Deionized water (1 ml) andchloroform (1 ml) were added to the solution causing it to separate into two phases. Thetop phase contained methanol, water, and the arsenicals while the bottom phase containedchloroform and the phospholipids.5.3.7 Phosphorus assayAn aliquot of the chloroform (15 .tl) from the Bligh-Dyer extraction was pipettedinto a 20 ml test-tube. Standards of 2 x i0 MNa2HPO4 (0, 50, 100 and 200 jil) werealso pipetted into separate 20 ml test-tubes. Perchioric acid (0.65 ml) was added to eachof the test-tubes which were then covered with a marble and placed in a metal test-tuberack which was sitting on a heating mantle which was turned to maximum heat. Afterabout 90 minutes the samples were allowed to cool and FISKE (0.75 ml) and ammoniummolybdate (7.0 ml) were added. The samples were vortexed and then placed into a steambath for 15 minutes. The standards were placed into a UV16OU Shimadzu spectrometerand a calibration curve was plotted. The unknown was placed into the spectrometer andthe phosphorus concentration was determined from the calibration curve.5.3.8 Processing the spectraA1H-NMR spectrum ofDMA both on the inside and the outside ofLUVs isdisplayed in Figure 5.7. The presaturated water suppression technique is effective inreducing/removing the water signal while not interfemng with the interesting region of thespectra. The large series of 4 peaks beween 3.3 and 2.6 ppm is from the Hepes buffer.124The broad peak centered at 1.78 ppm is the shifted and broadened (by Mn +2)spectroscopic signal of the methyl-group of the DMA molecules which are on the outsideof the LUVs. Likewise, the narrow peak at 1.65 ppm is the spectroscopic signal of theDMA on the inside oftheLUVs. The DMA and Hepes peaks were integrated as shown inFigure 5.7. The data were tabulated and the process was repeated for each of the 25spectra which constituted a diffusion experiment. An example of the raw data obtainedfrom one run is displayed in Table 5.1.1 . —2.6 2.4 2.2 2.0PP4* 1:4 12 10‘73:21EeFigure 5.7 The1H-NMR spectrum of DMA and LUVs.125Table 5.1 Raw data from a diffusion experimentExperimental thne (s) Hepes Integral DMA1 Integral DMA Integral0 304.09 45.22 29.10300 319.97 39.27 41.11600 310.79 35.63 45.35900 313.93 30.65 49.611200 309.56 27.70 50.771500 298.39 22.49 52.821800 304.34 22.67 54.242100 318.07 21.75 62.782400 294.20 17.89 57.152700 298.70 17.88 59.383000 302.97 17.26 61.813300 297.33 15.46 61.193900 301.98 14.65 62.834500 307.91 12.78 67.245100 303.49 12.22 65.295700 320.08 11.62 71.136900 316.10 12.28 70.668100 316.11 10.80 69.309300 293.88 7.00 66.1710500 300.51 7.24 68.1212300 313.33 9.24 71.5514100 300.45 7.23 66.8715900 305.51 6.88 70.3119500 323.43 9.21 72.9919800 319.66 7.81 72.40The integrals measured from the spectra at the various time intervals of the DMAmolecules on both the inside and the outside of the LUVs were divided by the Hepesintegral for calibration purposes. This process helped to minimize instrumental variabilityand the results of this process are displayed in Table 5.2 and in Figure 5.8. Instrumentalvariability is caused by changes to the shimming parameters and may be observed by thechanges to the Hepes integral during the experiment (Table 5.1).126Table 5.2 Integral ratios and time for a diffusion experimentExperimental time Integral ratio DMAj1/Hepes Integral ratio DMA0/Hepes(s)0 0.1487 0.0957300 0.1227 0.1285600 0.1146 0.1459900 0.0976 0.15801200 0.0895 0.16401500 0.0754 0.17701800 0.0745 0.17822100 0.0684 0.19742400 0.0608 0. 19422700 0.0599 0. 19863000 0.0570 0.20403300 0.0520 0.20583900 0.0485 0.20804500 0.0415 0.21845100 0.0403 0.21515700 0.0363 0.22226900 0.0388 0.22358100 0.0342 0.21929300 0.0238 0.225 110500 0.024 1 0.226712300 0.0295 0.228314100 0.0241 0.222615900 0.0225 0.230119500 0.0285 0.225719800 0.0244 0.2265The internal to external volume ratio was determined from the spectra that wereobtained when the system had reached equilibrium (ie. the spectroscopic signals wereneither increasing nor decreasing). The equilibrium position will be different forexperiments where glucose was used to balance the osmotic pressure than for theexperiments where NaCI was used to balance the osmotic pressure because of the salt-effect (see page 128).For the data in Table 5.2 the integral ratio for the outside and the inside DMAmolecules at equilibrium were determined to be 0.225 and 0.026, respectively. Theequilibrium concentrations of DMA is assumed to be equal on both sides of the membraneif glucose was used to balance the osmotic pressure, therefore, the value for f is written as:127+ + ++ +++(Jutside 4-4++ +0.2-++., O.l5ii÷+0.1:. ———O.05— —— Inside0— I I I I I0 2 4 6 8 10 12 14 16 18 20TIME(SECS.)(Thousands)Figure 5.8 An example of a plot of the inside (i and outside (4) integral ratios versustime for the difihsion of DMA through a liposomal membrane.128f Jeq/jeq0= vin/vout [5.56ajFor the data in Table 5.2; f 0.026/0.225 = 0.116.Since the total volume (Vtotaj) in the NMR tube is known:Vj + V01 =V0j= 0.6 ml [5.56b]Equations [5.56a] and [5.56bJ may be combined which allows V and V0to becalculated.Thus, V1 = 0.062 ml and V0= 0.538 mlWhen NaC1 is used to minimize the osmotic pressure, it is necessary to correct theequilibrium concentrations of DMA to account for the “salt-effect,” which is observedwhenever a large amount of inert electrolyte is added to a weak acid. The degree ofdissociation of the weak acid varies under these conditions. The “salt-effect” is increasedin magnitude as the concentration of the electrolyte is increased. The true value of f, forthe concentration ofNaC1 used in some of the difluision experiments is calculated asfollows:f 1apparent/(’Y±)2= 178apparent [5.57]where apparent is the integral ratio of the DMAin/DMAout at equilibrium in the presenceof NaC1 and (y) is the activity coefficient of the media in the presence of the NaC1.Equation [5.57] is derived as follows:The NaCI establishes a Gibbs-Donnan potential where;inside zGjnAG°jn+RTIn(Q)in [5.58]129outside iiGoutG°out+RTln(Q)out [5.59]where G (Joules or Calories) is the Gibbs free energy.At equilibrium, the chemical potential difference will be 0, therefore:zGm = AGot =0 [5.60]alSO, AG°m = AG°out [5.611therefore, Qin = Qout [5.62]and [MAiin[Hiin(Y±)2in/[JJin(Y±[DMAiout[Hjout(y±)2out/[DMA ] [5.63]however, (‘y±)jn 1 in the absence of any salt, for the [DMAH] both inside and outsideis also 1, and (Y±)out 1 because of the NaC1 on the outside of the LUVs. Therefore,in the presence of a salt:f = Ieqj/()2eq0= Ceqj/(y)2ceq0 [5.64]Equation [5.57] is derived for these experiments by substituting (y±) = 0.75 (the activitycoefficient for the concentration ofNaC1 used) into equation [5.60].The data in Table 5.2 is now plotted according to the linear equations:lfl(Ieq0-Itout) =- 10out) [5.67a]and ln(Itj - jeq = (1+f)ktfVjn+lfl(I0jn- Ieq0) [5.67b]or ln(Ieq0-=-10out) [5.68a]and ln(Itj - Ieq) = ..(1+f)k+ln(J°j - Ieq0) [5.68b]and these plots are displayed in Figure 5.9:130Figure 5.9 An example of a plot of in (nt..neq) () and In (+) versustime for the diffusion ofDMA through a liposomal membrane.TIME(SECS.)131The pH dependent rate constants (Ws) for the diffusion of the molecules, asmonitored by changes to the inside and the outside peak were determined from the slopes(-(1+f)k]V) or (-(l+f)k) of the lines.- The slope of the line for the inside peaks was -0.00056 ± 0.00003therefore, k = 0.000032 (± 0.000004) cm3 r1 and k = 0.00051 (± 0.00003) s_IThe slope of the line for the outside peak was -0.00062 ± 0.00005therefore, k = 0.00003 5 (± 0.000005) cm3 s’ and k = 0.00056 (± 0.00005) s_IThe rate constants for the inside and the outside sets of data which tended to agree withinabout 10% were averaged. Thus, k = 0.000034 (± 0.000009) cm3 r1 and ç = 0.00059 (±0.00010) s for the data from the experiment in Tables 5.1 and 5.2.The time (t) in the equations [5.47] and [5.48] will contains an t term which is thetime from the collection of the LUVs from the column to the time when the first FJD wasaquired. This term varied from 120 to 600 seconds and was timed by using a stopwatch.This time was added to each of the experimental times, therefore:ttexp+t [5.69]where t is the total time of diffusion from when the LUVs exited the column and texp isthe time from when the first spectra was recorded. Thus equations [5.67a and b] may berewritten as:ln(Ieq0- rout) = (1+f)ktexpNin+{l+f)ktNin + In(Ieq0- I°out) [5.70a]and ln(Itj - Ieq) = .(l+f)ktexpNin+{l+f)ktfVin + ‘flO0in -1out) [5.70b]The intercepts from the plots in Figure 5.9 should be equal and are written as:132Intercept = -(1+f)kt/Vj + ln(Ie%ut- T0out) [5.71a]= -(1+f)kt/Vj + ln(I°j - Ieq0 [5.71b]The values for the intercepts in Figure 5.9 were -2.09 ± 0.04 and -2.11 ± 0.06 forthe inside and the outside DMA integral ratios, respectively. t was measured to be 450seconds for the experiment in Figures 5.8 and 5.9. loin and ‘°out were then calculated tobe 0.189 ± 0.004 and 0.064 ± 0.002, respectively.The values for the terms were substituted into equations 5.47 and 5.48 and thecalculated values for the inside and outside DMA ratios were determined. The results aredisplayed in Table 5.3 and are plotted in addition to the experimental data (Table 5.2) inFigure 5.10.Table 5.3 Calculated integral ratios for the diffusion experiment in Table 5.2.Total time Calculated Integral ratio Calculated Integral ratio(s) DMA1/Hepes DMA,/Hepes450 0.1510 0.1015750 0.1307 0.12161050 0.1137 0.13831350 0.0995 0.15241650 0.0876 0.16421950 0.0776 0.17402250 0.0692 0.18232550 0.0622 0.18922850 0.0563 0.19503150 0.0514 0.19993450 0.0473 0.20403750 0.0438 0.20744350 0.0385 0.21264950 0.0348 0.2 1635550 0.0322 0.21896150 0.0303 0.22077350 0.0281 0.22298550 0.027 1 0.22409750 0.0265 0.224510950 0.0263 0.224812750 0.0261 0.224914550 0.0260 0.225016350 0.0260 0.225019950 0.0260 0.225020250 0.0260 0.2250133-1- 4- + +++ Outside0.2 +0.150J0.1. ±0.05—.— Inside0 I I I I I I0 2 4 6 8 10 12 14 20TIME(SECS.)(Thousands)Figure 5.10 An example of the inside (s) and outside (4) integral ratios versus time forthe diffusion of DMA through liposomal membranes. The solid lines arethe experimental fits for the data which were generated by using equations[5.47] and [5.48].1345.3.9 Permeability Coefficient DeterminationThe permeability coefficients were determined from the rate constants and thephosphorus assay results. The trapped volume (Vj) per mol of phospholipid (cm3i.tmo[1) was determined. The area of an EPC head group is estimated to be 60 Angstroms2,therefore, the area per iimol of phospholipid equals:= (6.O23xlO2molecules/mol x 106pmollmol x 60A2/lipid moleculex10-16cm2A)/2=1.81 x103cm2/i.t ol phospholipid [5.72]The permeability is determined according to the following:P = k(cm3s)V(cm3.tmol)/V(cm3A(2/pmol) [5. 73a]= k(s1)V(cm3imol)/A(cm2/ii.mol) [5. 73b]By using the phosphorus assaying technique, the total amount of lipid in the aboveexperiment was determined to be 0.0088 ± 0.00 10 mmols. Since the internal volume was0.062 cm3, the ratio of internal volume to lipid was 7.1 ± 0.9 cm3 mmol1. Therefore, thepermeability for the above experiment may be written as:P=(0.00059±0.00010r1x 7.1 ±0.9cm3s)I(1.81 x 103cm2/p. ol x jimoL/mmol)=2.3(±0.7)x 10 cms1355.4 RESULTS AND DISCUSSION5.4.1 pK determinations for MMA and DMAEither MMA or DMA was dissolved in deionized water and HCI was added to thesolution in small aliquots and the resulting changes in pH were monitored. The resultswere plotted and the pK’s were determined from the inflection points. The average resultsfor the pK’s from three potentiometric titrations are displayed in Table 5.3.Table 5.3 pK’s for MMA and DMApK pKMMA 4.1l(36)a 8.77(8.2)aDMA 6.28 (6.2)a -a) Literature values148 are given in brackets.The experimental value for DMA agrees with the literature values, however, theexperimental values for MMA seem to vary by 0.5 units of pK. (The MMA wassubmitted for micro-analysis and these results confirmed its purity.) The pH electrodewas standardized with fresh buffer at pH’s 4 and 7, therefore, the experimentally obtainedvalues were used in the calculations.In addition to the potentiometric titrations, the effects of pH upon thespectroscopic shift of the protons in the methyl group attached to the arsenic wasinvestigated (see top of page 120). Solutions of both MMA and DMA were made up atdifferent pH’s at the ionic strength of the diffusion experiments and the resonance138Table 5.5 The1H-NMR methyl resonance positions for MMA and DMA withvariations in pH.pH MMA resonance pH DMA resonanceposition (ppm) position (ppm)1.88 2. 1577 2.45 2.03963.49 2.0519 3.10 1.97214.19 1.9215 4.08 1.96475.16 1.8132 5.28 1.95615.39 1.8083 6.06 1.85725.65 1.8034 7.22 1.63347.43 1.7911 7.50 1.62168.16 1.7665 8.50 1.60538.56 1.7296 9.90 1.60009.02 1.6484 10.60 1.59779.83 1.5352 11.74 1.597710.22 1.505711.04 1.4885The variation in the methyl resonance position with pH allows the ipH inside ofthe LUVs to be monitored during an experiment. The buffering capacity on both theinside and the ouside of the LUVs was designed so that the ApH on both sides was 0.3pH units. The data in Table 5.6 and Figure 5.13 for the inside methyl resonance positionduring the diffusion experiments of DMA out of the LUVs at a pH 7.4 and a temperatureof 24°C shows that the pH on the inside of the LUVs is fairly constant and the ApH is0.3 pH units. The outside methyl resonance was more difficult to monitor because of thebroadening effects of the +2•Table 5.6 The1H-NIvIR methyl resonance positions for DMA during a diffusionexperiment.Experimental Time (s) Methyl resonance Experimental Time (s) Methyl resonanceposition ppm position ppm0 1.6287 4800 1.6189300 1.6262 5400 1.6213600 1.6189 6600 1.6164900 1.6213 7800 1.61391200 1.6139 9000 1.61641500 1.6164 10200 1.61641800 1.6213. 11400 1.61642100 1.6189 13200 1;61402400 1.6189 15000 1.61642700 1.6213 16800 1.61643000 1.6213 20400 1.61643600 1.6164 24000 1.6238136positions of the methyl group(s) at were plotted versus the pH. The tabulated data aredisplayed in Table 5.5 and the plots are displayed in Figures 5.11 and 5.12.2.22.12-1.91.81.71.61.51.41 ‘1 a a 11pH12Figure 5.11 A plot of the1H-NMR methyl resonance positions for MMA versus pH.2.05-2-1.95-1.91.85D 1.81.751.71.65-1.61.551372 epHl’i 12Figure 5.12 A plot of the1H-NMR methyl resonance positions for DMA versus pH.1391.71.681.661.64r-.1.62 ——— — _ —— R — —1.61.56-1.56-1.54-0 5 10 15 25Time (sec)(Thousands)Figure 5.13 A plot of the1H4.44R methyl resonance positions for DMA versus timefor a diffusion experiment.1405.4.2 The diffusion of DMA through LUVs of various sizesDMA was encapsulated by EPC and the mixture was freeze-thawed, extrudedthrough filters and passed down a Sephadex column as described in Section 5.3.4. Thepore sizes of the filters (50, 100, 200, and 400nm) were varied to produce the desired sizeofLUVs. Glucose was used to balance the osmotic pressure and each of the experimentswas performed at a pH of 7.4 and a temperature of 24°C. The values listed in Table 5.7for Ic, k, and P are the pH dependent rate constants and permeability coefficients.Table 5.7 DMA diffusion through various sized LUVs (Temp = 24°C andpH=7.4).filter pore size average size of Ic (s1) x k (cm3s1)x P (cm s1) x(nm) LUVs (nm)’49 io.950 68±19 3.8±0.6 2.7±0.7 1.6±0.5100 103±20 3.4±0.5 2.9±0.7 1.6±0.5200 151 ± 36 2.9 ± 0.4 2.7 ± 0.7 1.5 ± 0.5400 243 ± 91 1.4 ± 0.2 1.3 ± 0.3 0.6 ± 0.2The values for the rate constant k decrease as the size of the LUVs increase. Thisis expected as its magnitude should vary with iIr, where r is the radius of the vesicle (seeSection 5.2.6). Incorporating the internal volume into the rate constant to obtain thevalues for k produces rate constants which are more independent ofvesicle size. Indeed,this is certainly the case for LUVs produced by extrusion through filters of 50, 100, and200 mn pore sizes. These values are expected to vary with the area of the vesicles (see141Section 5.2.6), however, the total surface area of the vesicles in the first three experimentswill be approximately the same. This is because the same amount of lipid was used in eachcase and the phospholipids arrange themselves into a single bilayer. When filters of400nm pore size are used the phospholipids assemble in multiple bilayers. It has beenshown that only 30% of the phospholipid is on the outside, while 70% remains on theinside of the vesicles149. This leads to decreased total surface area which is why k issmallest for the LUVs produced by extrusion through the largest pore size filters. Thepermeability coefficients are theoretically independent of vesicle size (see Section 5.2.6);however, as it is defined as DKJAx, the permeability coefficient should decrease as thethickness of the bilayer Ax increases. The LUVs produced which have a single bilayer allhave similiar permeability coefficients, while the LUVs produced which contain somemultiple bilayers have a decreased permeability relative to the others.5.4.3 DMA pH studyDMA was encapsulated by EPC and the mixture was freeze-thawed, extrudedthrough 200 nm filters, and passed down a Sephadex column as described in Section5.3.4. Both Glucose and NaCl were used to balance the osmotic pressure and theexperiments were performed at 5 different pHs at a temperature of 24°C. The valueslisted in Tables 5.8 and 5.9 for Ic, k, and P are the pH dependent rate constants andpermeability coefficients, respectively, and are the average values obtained afterperforming the experiments in duplicate. The data for the rate constants was plotted vsthe [Hj (Figures 5.14 and 5.15).142Table 5.8 DMA pH Study (Temp = 24°C with NaC1 outside)pH Ic (s’) x104 k (cm3s1)x105 P (cm s’) x1097.00 11.4± 1.7 14.5 ±3.6 11.0 ±3.37.15 8.5±1.3 8.9±2.2 3.9±1.27.40 6.0±0.9 5.7±1.4 3.0±0.97.73 2.3 ± 0.3 2.4 ± 0.6 1.5 ± 0.57.97 1.9±0.3 1.6±0.4 1.4±0.4Table 5.9 DMA pH Study (Temp = 24°C with glucose outside)pH Ic (s-i) x104 k (cm3r1)x105 P (cm s’) x1097.00 18.5±2.8 12.8±3.2 8.5±2.57.15 6.8±1.0 8.1±2.0 4.4±1.37.40 5.8 ± 0.8 5.2 ± 1.3 2.8 ± 0.87.73 2.9±0.4 3.0±0.8 1.5±0.57.97 1.50 ± 0.2 1.6 ± 0.4 1.2 ± 0.4The slope and the intercept for the plot in Figure 5.14 are 1300±68and (1.27 ± 7.18) x i06, respectively, while the slope and intercept for the plot in Figure5.15 are 1400 ± 200 and (-7.13 ± 220) x 10-6, respectively. These results for the efflux ofDMA provide conclusive evidence that the neutral form of the molecule is dominating thediffusion. As the [Hj approaches 0 the percentage of DMA- approaches 100. Thishypothetical situation is143E0IFigure 5.14 A plot of the pH dependant rate constants (k in cm3s-i) vs the [Hi for theefflux ofDMA at 24°C. Either glucose (a) or NaC1 (ii) was added to theoutside of the 200 nm LUVs to balance the osmotic pressure.1412100 0.1 0.2 0.3 0.5 0.6[11t]/0.7 0.8 0.9144Figure 5.15 A plot of the pH dependant rate constants (jç in s-i) vs the [H+J for theefflux ofDMA at 24°C. Either glucose (a) or NaCI (i)was added to theoutside of the 200 nm LUVs to balance the osmotic pressure.0.5[H’/l 0-7(Times 1 OE-7)w145represented by the y-intercepts on the graphs in Figures 5.14 and 5.15 which are both 0within the uncertainty of the line. Therefore, the rate constants for DMA- are notmeasurable by these experiments and are at least 1 to 1 O times slower than the rateconstants for DMAH. This is a conservative estimate based upon equation [5.43] becausethe overall rate constant is a combination of the individual rate constants times theconcentration of each molecular species. The concentration of the anion is 10 to 100 timesthat of the neutral species at the pH’s investigated and it still does not make a contributionto the overall rate constant, hence the rate constant for the anionic species must be verysmall.By using equations 5.50a, b, and c the values for k&jj, kaji, and Ppj.j at 24°Cthrough 200 nm LUVs were determined to be 9.28 (± 0.56) x i0 cm3r’, 10.1 (± 1.4) x10-3 s-1, and 5.8 (± 1.0) x 10-8 cm s, respectively. These values are independent ofpH.There is a much higher percentage of error on the slopes and the intercepts for thegraph in Figure 5.15 than the graph in Figure 5.14. This indicates that incorporating thetrapped volume into the rate constants decreases the variability between experiments andproduces results which are more reproducible.These experiments also demonstrate that either NaCI or glucose may be used onthe outside of the LUVs to balance the osmotic pressure because when the equilibriumposition is corrected for the salt effect by using equation 5.57 the results displayed inTables 5.8 and 5.9 are approximately equal.5.4.4 MMA pH and temperature studyMMA was encapsulated by EPC and the mixture was freeze-thawed, extrudedthrough 200 nm filters, and passed down a Sephadex column as described in Section5.3.4. NaC1 was used to balance the osmotic pressure and the experiments were146performed at 5 different temperatures and 2 different pHs. The MMA diffused out of theLUVs very slowly and N14R spectra were taken 3 times a day for about a week. Thesamples were placed in water baths at the appropriate temperatures when spectra were notbeing recorded. The results for these experiments are displayed in Tables 5.10 and 5.11.Table 5.10 MMA Diffusion Coefficients (DH = 6.98) through 200 urn LUVs.Temp (°C) k (cm3r1)x 106 ç (r’) x P (cm s’) x 10-1124.0 0.53 ±0.13 0.27±0.04 0.69±0.2129.4 0.69 ± 0.17 0.36 ± 0.05 0.88 ± 0.2631.0 1.37 ± 0.34 1.02 ± 0.15 1.31 ± 0.3938.5 2.06 ± 0.52 1.06 ± 0.15 2.63 ± 0.7947.8 12.5±3.1 9.3±1.4 11.9±3.6Table 5.11 MMA Diffusion Coefficients (pH = 7.40) through 200 urn LUVs.Temp (°C) k (cm3rl) x i06 Ic (s1) x P (cm r1) x 10-1124.0 0.20 ± 0.05 0.11 ± 0.02 0.28 ± 0.0829.4 0.46±0.12 0.26±0.04 0.63 ±0.1931.0 0.38 ± 0.09 0.33 ± 0.05 0.70 ± 0.2138.5 1.46±0.37 0.81 ±0.12 2.01 ±0.6047.8 5.3±1.3 4.6±0.7 9.8±2.9147At a pH of 6.98 the species and percentages of each are; MMAH (0.133%),MMA- (98.3%) and MMA-2(1.59%) while at a pH of 7.40 the species and percentagesof each are; MMAH (0.049%), MMA- (95.8%) and MMA2(4.08%). The pHindependent rate constants and permeability coefficients are calculated according toequations 5.50a, b, and c and the results are displayed in Tables 5.12 and 5.13. Thediffusion coefficients in Table 5.12 are from the experiments at pH 6.98; while thediffusion coefficients in Table 5.13 are from the experiments at pH 7.4.Table 5.12 MMAH Diffusion Coefficients through 200 nm LUVs (from Table 5.10.Temp (°C) k.L&J{ (cm3r1) 1c4&j.j (sd) PAJ] (cm s)x 10-6 x i- x i01124.0 400± 120 205 ±50 780±27029.4 520± 160 270±70 1020±36031.0 1040±310 770± 190 2580±90038.5 1550±470 800±200 3030±106047.8 9400 ±2800 7000± 1700 23500±8300Table 5.13 MMAH Diffusion Coefficients through 200 nm LUVs (from Table 5.1 1).Temp (°C) kp.j (cm3r1) icpj.j (s-i) P&jj (cm s-i)x 10-6 io-5 x 10-il24.0 410±120 230±60 880±31029.4 930±280 520±130 1980±69031.0 760±230 670±170 2310±69038.5 2960±890 1660±420 6320±220047.8 10700±3200 9390±2300 32500± 11000148These results indicate that the neutral form of the molecule is dominating thediffusion, because the pH independent diffusion coefficients calculated from the pHdependent diffusion coefficients at pHs 6.98 and 7.40 are relatively close.Arrhenius plots of the pH independent permeabilities for MMA (from the diffusiondata obtained at pHs 6.98 and 7.4) versus the inverse temperature are plotted according toequation 4.8 and are shown in Figure 5.16. The slopes of these lines equal -14300 ± 700and -13200 ± 2400 from the diffusion experiments at 6.98 and 7.4, respectively. TheFigure 5.16 A plot of the ln(PL&J{) from the diffusion experiments at pHs 6.98 ( ) and7.4( )versus l/T.149average activation energy for the permeation of the MMAH through the phospholipidbilayer was calculated by using equation 4.8 and is displayed in Table 5.14.Table 5.14 The activation energy for the permeation ofMMAH through thephospholipid bilayer of LUVs.Compound Activation EnergyMMA}1 114±lOKJmolThese experiments were performed at pHs where the percentage of the anionicspecies was approximately 1000 times greater than that of the neutral species and,therefore, an attempt was made to determine the diffusion coefficients for the anionicspecies.At pH 6.98, Pp.j(O.OOl33)+A(0•983)=P698likewise at pH 7.4 Paj(0.00049) +A(0•958 =Thus, at each temperature there are two unknowns and two equations. When theequations are solved the results which are presented in Table 5.15 are obtained. Theerrors on the values for A- are estimated to be at least 200% and, therefore, are nottabulated.Table 5.15 Permeability coefficient determination for MMA.Temp (°C) PA}I (cm s-i) PA-(cmr1)x i013 iO-1324.0 47400 4.8729.4 28600 51.131.0 72100 36.038.5 69200 17447.8 225000 908150If a similar type of reasoning is applied to the rate constants the results displayed in Table5.16 are obtained. The errors on the values for A- are estimated to be at least 200% and,therefore, are not tabulated.Table 5.16 Rate constant determination for MMA.Temp (°C) kj (cm3s1) kA- (cm3s4) icpLJ.J (s-i) icA- (s-i)x iO-8 x 10-8 x io-7 x24.0 39000 1.14 19000 2.0829.4 26800 33.9 11300 20.831.0 119500 -22.0 82500 -7.9038.5 68300 117 27200 71.147.8 854000 113 546000 203The scatter of these results and a few negative numbers indicates that the error inthese numbers is very high. However, these results show that the diffusion coefficients forthe anionic form of the molecule seem to be approximately 1000 to 10000 times smallerthan those for the neutral form of the molecule.5.45 DMA temperature studyDMA was encapsulated by EPC and the mixture was freeze-thawed, extmdedthrough 200 nm filters, and passed down a Sephadex column as described in Section5.3.4. These experiments were performed at 4 different temperatures all at a pH of 7.4.Glucose was used to balance the osmotic pressure. The results from these experimentsare displayed in Table 5.17.151Table 5.17 DMA Diffusion Coefficients (pH 7.4) through 200 nm LUVs.Temp (°C) k (cm3s1) ic (s-i) P (cm s-i)x i- x i- x24.0 5.2±1.3 5.8±0.9 2.8±0.828.0 6.3±1.6 9.9±1.5 3.8±1.130.0 8.0 ± 2.0 7.0 ± 1.0 4.1 ± 1.233.0 10.3±2.6 15.6±2.3 8.3±2.5The pH independent values for these experiments are calculated according toequations 5.50a, b, and c and the results of these calculations are displayed in Table 5.18.Table 5.18 DMAH Diffusion Coefficients through 200 nm LUVs.Temp (°C) kp.j.j (cm3s) (r1) Pp.j (cm s)x iO-5 x x24.0 87±22 97± 15 46± 1428.0 107±26 166±25 63±1930.0 135±34 118±18 68±2033.0 173 ±43 263 ±40 139±41An Arrhenius plot of the pH independent permeabilities versus the inversetemperature according to equation 4.8 is displayed in Figure 5.17. The slope of this lineequals -10400 ± 2800, while the intercept equals 18.0 ± 0.2. The activation energy for the152permeation of the DMAH through the phospholipid bilayer was calculated by usingequation 4.8 and is displayed in Table 5.19.Table 5.19 The activation energy for the permeation ofDMAH through thephospholipid bilaver ofLUVs.Compound Activation EnergyDMAH 86±20 KJmol’-15.9--16--16.1--16.2--16.3--16.4--16.5--16.6--16.7--16.8-16.9--17-0.00326Figure 5.17 A plot of the ln(P&j) versus l/T for the activation energy determinationfor DMAH diffusion through a phospholipid bilayer.0.00328 0.0033 0.00332 0.00334 0.003361/T(K)153The activation energy for the diffusion ofMMAH across the liposomal membranes(114 KJ moll) is greater than the activation energy for DMAH (86 KJ mol1), asdetermined by using the NMR technique. A similar result was observed earlier in Section4.4.2, although the difference in the activation energies for MMA and DMA is less (MMAwas found to have an activation energy of 220 KJ mo11 and DMA was found to have anactivation energy of 130 KJ mol1 by using the radioactive method and 100 nm LUVs).MMA has a greater hydrogen bonding capability with the surrounding water moleculesthan does DMA and, therefore, would be expected to have a greater activation energy.Indeed, it has been shown in previous studies with erythritol, glycerol, and glycol15°thatthe activation energy for permeation through artificial membranes corresponds directly tothe number of hydrogen bonds that need to be broken in order for the permeant toundergo complete dehydration.5.4.6 DMA temperature study with cholesterol in the membraneDMA was encapsulated by EPC and the mixture was freeze-thawed, extrudedthrough 200 nm filters, and passed down a Sephadex column as described in Section5.3.4. These experiments were performed at a pH of 7.0 with 5 different temperatures.Glucose was used to balance the osmotic pressure. The results from these experimentsare displayed in Table 5.20.The pH independent values for these experiments are calculated according toequations 5.50a, b, and c, and the results of these calculations are displayed in Table 5.21.154Table 5.20 DMA Diffusion Coefficients (pH 7.0) through 200 nm LUVs (55:45,EPC :Cholesterofl.Temp (°C) k (cm3r1) k (s1) P (cm s)x iO-5 x x 10-1022.0 1.6±0.4 1.3±0.2 3.8±1.127.0 2.8±0.7 3.1±0.5 7.8±2.332.0 4.5±1.1 4.8±0.7 10.9±3.337.0 11.2±2.8 11.1± 1.7 20.7±6.242.0 14.2±3.6 14.8±2.2 29.6±8.9Table 5.21 DMAH Diffusion Coefficients through 200 nm LUVs (55:45,EPC:Cholesterol).Temp (°C) kpj.j (cm3rl) i.cpj (s-i) Pp.j1 (cm s)io-5 x 10-’ x 10-1022.0 11.7±3.0 9.1±1.4 28±827.0 20.3±5.1 22.7±3.4 57±1732.0 33.0±8.3 35.0±5.3 80±2437.0 82±21 80± 12 151 ±4542.0 104±26 107±16 216±65An Arrhenius plot is constructed, Figure 5.18, of the pH independentpermeabilities versus the inverse temperature according to equation 4.8. The slope of thisline equals -9440 ± 580, while the intercept equals 12.3 ± 0.1. The activation energy for155the permeation of the DMAH through the phospholipid bilayer containing 45 mol %cholesterol was calculated by using equation 4.8 and is displayed in Table 5.22.Figure 5.18 A plot of the In(PMj) versus l/T for the activation energy determinationfor DMAH diffusion through a phospholipid bilayer which containscholesterol.-17.5•-18--18.5--1 9-19.5--20-0.00315.0.0032 0.00325 0.0033 0.00335irr0.003156Table 5.22 The activation energy for the permeation ofDMAH through a phospholipidbilayer which contains cholesterol.Compound Activation EnergyDMAH 78±5KJmo[’The addition of cholesterol to the membrane decreases both the rate constants andthe permeability coefficients for DMA (see Tables 5.17, 5.18, 5.20, and 5.21). This isexpected as the addition of cholesterol to the phospholipids will decrease the viscosity ofthe hydrocarbon region of the membrane,’5’which will result in the decrease of thediffusion coefficients for DMA. Addition evidence which supports a decrease in theentropy of the LUVs following the addition of cholesterol is provided by the Arrheniusplot in Figure (5.18). The activation energy (78 KJ moll) for when cholesterol isincorporated into the membrane is approximately the same (within experimentaluncertainty) as the activation energy (86 KJ moP1)when the membrane is composed onlyof phospholipid. However, the Arrhenius constants which relate to the entropy of themembrane are 6.6 x i07 and 2.2 x for the phospholipid and thephospholipid/cholesterol membranes, respectively. This result is consistant with thefindings of De Gier et a!. which showed that although the permeability coefficients maychange as the composition of the bilayer changes, the activation energy for a permeant willbe independent of the bilayer composition.’5°1575.4.6 Comments on how the results obtained in this chapter apply to biologicalsamples and possible uses for the technique in the future.The results in this chapter show that DMA will pass through biological membranesmuch more rapidly than MMA, if the only mechanism for transport through themembranes is via slow passive diffi.ision. The neutral species of each of these weak acidsis the major species which permeates through the membrane and the rate constant andpermeability coefficients at 24°C for the neutral DMA are 1.08 (± 0.08) x i0 cm3s1and 3.3 (± 0.08) x i08 cm s1. The rate constant and permeability coefficient at 24°C forthe neutral MMA are 3.93 (± 0.05) x 10 cm3r’ and 5.44 (± 0.05) x cm s-’,respectively. Thus, there is a pH dependence on the diffusion coefficients and as the pHdecreases the corresponding diffusion coefficients increase in magnitude. A culture ofC.humicola was innoculated with DMA and produced 87 nmoles of trimethylarsine at pH5 while only 41 and 2 nmoles of trimethylarsine were produced at pHs 6 and 7,respectively. 152,153 Under identical conditions, a culture inoculated with MMAproduced 9, 6, and 0 nmols of trimethylarsine at pHs 5, 6 and 7, respectively. Assumingthat the rate limiting step is the diffusion of the DMA or MMA into the cell, then theseresults may be explained by higher diffi.ision coefficients with decreased pH. Also, there ismuch more trimethylarsine produced for the experiments when DMA is the startingsubstrate rather than MMA. This is similiar to the results reported in Chapter 3 whichshowed that arsenobetaine is produced more readily when DMA is the precursor ratherthan MMA. It may be argued that the differences in the diffusion coefficients ofDMA andMMA are the main reasons for the differences in these two experiments. The compositionof the membrane effects the diffusion coefficients ofDMA and MMA as it was shown thatthe addition of cholesterol to the membranes decreases the diffusion coefficients.Ideally, the NMR techniques to determine diffusion and permeability described inthis chapter would be applicable to a wide range of molecules. However, it is limited to158molecules which have pH dependent permeability coefficients of lx i08 cmotherwise the molecule permeates too quickly to be measured. The technique could beused to investigate the permeation of other environmentally sensitive compounds and theeffects of these compounds on membranes. Perhaps the most useful application willinvolve the use of DMA as a probe molecule to study the effects of altering the membraneproperties upon the diffusion coefficients. The composition of the membrane could bealtered by using different combinations of phospholipids, by adding various amounts ofcholesterol and perhaps even some ionophores such as valinomycin.5.5 SUMMARYThe work presented in this thesis provides information about thebiotransformations and biomobiity of MMA and DMA in the marine environment. Thiswork shows that both MMA and DMA are metabolic precursors to arsenobetaine and thatmicroscopic organisms in the water column are responsible for these biotransformations.Mussels do not appear to be capable of synthesizing arsenobetaine which isbioaccumulated by mussels from seawater. DMA is more readily biotransformed toarsenobetaine than is MMA. This difference is postulated to be the result of different ratesof diffusion of these compounds into cells and this notion is supported by the results fromdiffusion experiments. Initially, these experiments were carried out with radiolabeledpermeants and model cells (liposomes) and were later modified by using ‘H-NMRmethodology in conjunction with shift agents which enabled the spectroscopic signal ofthe permeant on both the inside and the outside of the liposomes.159CHAPTER 6THE SYNTHESIS OF ORGANOARSENICALS RELEVANT TO THIS THESIS6.1 INTRODUCTIONMuch of the work in the preceeding chapters involved arsenic compounds whichwere not readily available. The synthesis of the[3H1-MMA and[3H]-DMA was essentialfor the studies on the biotransformations in chapters 2 and 3, and on the biomobility inchapter. The synthesis of arsenobetaine was very important because its identification inseawater and mussels was based upon chromatographic comparisons with the syntheticstandard. In addition, trimethylarsine, trimethylarsine oxide, and tetramethylarsonium ionwere synthesized. The details of these syntheses are contained in this chapter.6.2 EXPERIMENTAL AND RESULTS6.2.1 Synthesis of[3H1-MMA from arseniteThe following procedure is based upon a method outlined in reference 162, butchanges were made in order to incorporate the[3H]-label. Arsenite (2.8 g) was dissolvedin 10 M NaOH (8 ml) and the solution was added to a Carius tube (25 ml). The[3H1-.methyl iodide (10 mCi) in 1 ml of toluene was obtained from Amersham, USA in a smallcylindrical, sealed glass vial fitted with a break seal. The break seal was smashed with aspatula and the contents of the vial were poured into the reaction tube along with thedeionized water (1 ml) which was used to rinse out the vial. Unlabelled methyl iodide (2ml) was also added to the reaction tube which was sealed and heated at 120°C with a sandbath. After 48 hours the reaction tube was removed from the sand bath and white crystals160were precipitating out of the solution. The reaction tube was cooled in an ice bath andadditional crystals precipitated out of solution. The reaction tube was cracked open andthe solution was poured out leaving the crystals in the tube. Ethanol (2 x 10 ml) wasadded to the solution to precipitate the remaining crystals (5.898 g) which were removedby filtration. The crystals were identified as the Na salt of[3HJ-MMA and the purity wasexamined by using ‘H NMR spectroscopy. The1H-N?i6R (200 MHz in D20) spectrum,Figure 6.1, shows a singlet at 1.48 ppm. This corresponds with the resonance position forthe singlet found in the literature for the ‘H-NMR spectrum for MMA (at pH 12.6 inD20).163BRAP300S. 120AU PROG:X0D.AUDATE 30—4—90SF 200.133SY 90.001 3950.000SI 32758TO 32768SW 4000.000HZ/PT .244PW 0.0RD 0.0AD 4.096RD 8MS 32TE 298FM 500002 3630.000OP 63L PDLB .300GB .100CX 25.00CY 17.00Fl 1l.500PF2—1.000PHZ/CM 100.066PPM/CM .500SR 2834.0311.0 10.0 9.0 8.0 7.0 6.0 5.0 4.0 3.0 2.0 1.0 0.0PPMFigure 6.1 1H-NMR (200 MHz in D20) spectrum of[3H1-MMA.1616.2.2 Synthesis of[3H]-DMA from[311]-M.MA[3H]-MMA (2.0 g) was dissolved in the minimum amount ofwarm deionizedwater ( 10 cm3), then sulphur dioxide was bubbled through the solution until saturationoccured. The solution was boiled for 2 minutes, quickly cooled to 4°C, and neutralizedwith sodium carbonate. The solution was evaporated to dryness and the methylarsineoxide was extracted from the residue by using benzene. Removal of the benzene left afoul-smelling solid. This was dissolved in the minimum amount of methanol ( 15 cm3),and placed in a Carius tube. Methyl iodide and sodium hydroxide slightly in excess ofstoichiometric amounts were added and the reaction tube was sealed and heated at 45°Cfor three days. The methanol was then evaporated and the residue was redissolved in aminimum amount of deionized water. Hydrogen peroxide (0.6 cm3, 30%) was slowlyadded and the excess was boiled off The reaction mixture was then added to a SephadexLH-20 column (1 cm x 30 cm) and eluted with water. The first 60 cm3 was collected andthe water was evaporated. The sample was then added to an Amberlite IRA-410 anionexchange column (1.5 x 70 cm) and eluted with water. The arsenic compound (0.56 g)eluting between 60 and 190 cm3 was isolated as the Na salt of[3}{1-DMA. This wasconfirmed by using 1H NMR spectroscopy. The1H-NMR (200 MHz in D20) spectrum,Figure 6.2, shows a singlet at 1.60 ppm. This corresponds with the resonance position forthe singlet found in the literature for the1H-NMR spectrum for DMA (at pH 12.6 inD20).1631620 ABRSEI3OS. lOBAU PROS:X00.AUDATE 13—9—90SF 200.133S’( 80.001 3950.000SI 32768TD 32768SW 4000.000HZ/PT .244PW 0.0RD 0.0AD 4.096PG 10NS 32TE 298FM 500002 3530.000OP 63L. P0LB .30068 .100CX 25.00CY 17.00Fl li.SOOPF2 —i.000PHZ/CM 100.068PPM/CM .500SR 2834.03iflr - —11.0 10.0 9.0 0.0 7.0 6.0 4.0 3.0 2.0 1.0 0.0Figure 6.2 1H-NIvIR (200 MHz in D20) spectrum of[3H1-DMA.6.2.3 Synthesis of trimethylarsine from arsenic trichiondeThe following proceedure is based upon a synthesis outlined in reference 164. Atwo necked (250 ml) round bottomed flask was equipped with a reflux condensQr, a•stirrer, a separatory funnel (which contained methyl iodide (10 ml) in dry di-n-butyl ether(10 ml)) and a Tjoint so that a positive pressure of argon could be maintained through out163the reaction. Magnesium turnings (4.02 g), dry di-n-butyl ether (20 ml) and a few crystalsof 12 were added to the flask. The Grignard reaction was initiated by adding a few dropsof the contents of the separatory funnel to the stirred magnesium/dibutyl ether mixture.AfIer the reaction had started the remaining methyl iodide solution was added dropwiseover the period of 1 hour. The flask was cooled with an ice/water bath during theaddition. After all the methyl iodide had been added, the cloudy mixture was gentlyheated for approximately 30 minutes and then recooled by using the ice/water bath. Asmall amount of the magnesium remained unreacted. Arsenic trichloride (10 g in 25 ml ofdry di-n-butyl ether) was added dropwise through the addition funnel to the cooledreaction mixture (ice/water bath). After the addition of the arsenic trichloride solution(which took about 20 minutes) the reaction mixture was stirred for approximately 2 hoursat room temperature. The condenser was then replaced with a distillation apparatus. Themixture was heated to decompose the (CH3)AsMgX2adduct. Trimethylarsine andsome di-n-butyl ether distilled under these conditions and were collected and stored in atube that was sealed with a Teflon screw type apparatus. The mixture was redistilled toisolate trimethylarsine which was divided up and stored in sealed glass tubes.Trimethylarsine is a clear liquid and is very poisonous and flammable upon contact withair. Its synthesis was confirmed by its use in subsequent reactions (Sections 6.2.4, 6.2.5and 6.2.6).6.2.4 Synthesis of tetramethylarsonium iodide from trimethylarsineA reaction tube containing methyl iodide (1.42 g) in dry diethyl ether (4 ml) wasconnected to a vacuum line and cooled in liquid nitrogen. A tube containingtrimethylarsine (0.306 g) was also connected to the vacuum line and the contents weretransferred under vacuum to the reaction tube. The reaction tube was sealed and then164warmed to room temperature. White crystals started to form almost immediately and thereaction was allowed to proceed for approximately 4 hours. After the reaction had beencompleted the crystals which were identified as tetramethylarsonium iodide were filteredfrom the solution and air dried. The purity of the crystals (0.135 g) was checked by using‘H NMR spectroscopy and DCI-MS. The1H-NMR (200 MHz inD20) spectrum, Figure6.3, shows a singlet at 1.95 ppm. This corresponds with the resonance position for thesinglet found in the literature for the ‘H-NMR spectrum for tetramethylarsoniumiodide.99 The DCI mass spectrum is displayed in Figure 6.4. The base peak correspondsto the tetramethylarsonium ion.BRAP300S. 120AU PROS:X00.AUDATE 30—4—90SF 200.133SY 80.001 3950.000SI 32768TO 32768SW 4000.000HZ/PT .244PW 0.0RD 0.0AD 4.096RG 8MS 32it 298FM 500002 3630.000OP 63L PDLB .300GB .100cx 25.00CV 17.00Fl 11.500PF2 —1.000PHZ/CM 100.068PPM/CM .500SR 2834.0311.0 10.0 9.0 8.0 7.0 6.0 5.0 4.0 3.0 2.0 1.0 0.0PPMFigure 6.3 ‘H-NMR (200 MHz in 1)20) spectrum of tetramethylarsonium iodide.1651ee 12113575I8z hzzs,_ 92185CA.r ,ii Ii, ,.11 II —_....4 .—- . .-.— . .—, •148 68 88 188 128 148 168 188 288 228 248 268 m/zFigure 6.4 DCI mass spectrum of tetramethylarsonium iodide.6.2.5 Synthesis of trimethylarsine oxide from trimethylarsineA reaction tube containing hydrogen peroxide (1.5 ml) in dry diethyl ether (7 ml)was connected to a vacuum line and cooled in liquid nitrogen. A tube containingtrimethylarsine (0.642 g) was also connected to the vacuum line and the contents weretransferred under vacuum to the reaction tube. The reaction tube was sealed and thenwarmed to room temperature. The ether layer was initially cloudy, but became clearer asthe reaction proceeded. There was a continual stream ofbubbles from the aqueous to theether layer. After 24 hours the contents of the reaction tube were transferred to a roundbottom flask and the ether and water was rotary evaporated off leaving a white lava-likesolid. This solid was quickly sublimed at one atmosphere and stored in a sealed vial whichT166was placed into a dessicator. The purity of the crystals (0.198 g) was checked by using‘H NMR spectroscopy and DCI-MS. The ‘H-NMR (200 MHz in D20) spectrum, Figure6.5, shows a singlet at 1.7 ppm. This corresponds with the resonance position for thesinglet found in the literature for the1H-NMR spectrum for trimethylarsine oxide.165The DCI mass spectrum is displayed in Figure 6.6. The base peak of the spectrum oftrimethylarsine oxide corresponds to [M + Hj ions.BRMA28OS.123AU PROG:X00.AUDATE 28—5—91SF 200.133SY 80.001 3950.000SI 32768TD 32768SW 4000.000HZ/PT .244PM 0.0RD 0.0AG 4.096RG 16MS 32TE 298FM 500002 3630.000OP 63L PaLB .300GB .100CX 25.00CV 17.00Fl 11.500PF2 —i.000PHZ/CM 100.068PPM/CM .500SR 2834.03-IC,z11.0 10.0 9.0 8.0 7.0 6.0 5.0 4.0 3.0 2.0 1.0 0.0PPMFigure 6.5 1H-NMR (200 MHz in D20) spectrum of trimethylarsine oxide.167188x 13775-,56Z511*1.111 p_lt.I I I•I I I - I I I’I I46 66 88 168 128 148 168 188 288 228 246 266 mJzFigure 6.6 DCI mass spectrum of trimethylarsine oxide.6.2.6 Synthesis of arsenobetaine from trimethylarsineThe following proceedure is based upon a synthesis outlined in reference 164. Areaction tube containing ethyl-bromoacetate (1.5 g) in dry dibutyl ether (4 ml) wasconnected to a vacuum line and cooled in liquid nitrogen. A tube containingtrimethylarsine (0.45 g) was also connected to the vacuum line and the contents weretransferred under vacuum to the reaction tube. The reaction tube was sealed and thenwarmed to room temperature. The clear reaction became cloudy and white crystalsstarted to precipitate out of solution. After 24 hours the solvent was decanted and thewhite crystals were dissolved in a minimum amount of deionized water. This was appliedto an Amberlite IRA-410 column (70 x 1.5 cm in the OH- form) and was eluted by using168water. Aqueous eluant (150 ml) was collected and rotary evaporated to dryness leavingwhite crystals which were identified as arsenobetaine and were dried on the vacuum line.The purity of the crystals (0.353g) was checked by using 1H NMR spectroscopy and DCI-MS. The1H-NMR (200 MHz inD20) spectrum, Figure 6.7, shows a singlet at 2 ppmfrom the 9 identical protons attached to the methyl groups and a singlet at 3.5 ppm fromthe 2 identical protons in the methylene group. This corresponds to the1H-NMRspectrum found in the literature for arsenobetaine.43 The ratios of the areas is 4.52 whichcorresponds to the expected value of 4.5. The DCI mass spectrum is displayed in Figure6.8. This spectrum indicates a protonated molecule as well as a fragmentation pattern thatallows for positive identification of the compound.BRBJY180S.108AU PROG:X00 . AUDATE 18—7—91SF 200.133SY 80.001 3950.000SI 32768ID 32768SW 4000.000HZ/PT .244PM 0.0RD 0.0AG 4.096RG 40NS 32TE 298FM 500002 3630.000OP 63L P0LB .300GB .100CX 25.00CV 17.00Fl ii.500PF2 —2.000PHZ/CM 100.068PPM/CM .500SR 2834.03-. 1 .. I • 1•.1• ..11.0 10.0 9.0 8.0 7.0 6.0 5.0 4.0 3.0 2.0 1.0 0.0Figure 6.7 ‘H-NMR (200 MHz inD20) spectrum of arsenobetaine.56ILII16975(a,fl1J92Z5(AsrjtK4H146 66 88 188 128 148 166 186 288 2.28 248 268 m/zFigure 6.8 DCI mass spectrum of arsenobetaine.170BIBLIOGRAPHY1. National Research Council, Arsenic; National Academy of Sciences: WashingtonDC, 1977, p16.2. Sanders, 3G. Marine Em’. 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The biotransformations and biomobility of monomethlarsonate and dimethylarsinate in the marine environment
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