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Arsenic and antimony species in the terrestrial environment Koch, Iris 1998

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ARSENIC AND ANTIMONY SPECIES IN THE TERRESTRIAL ENVIRONMENT by IRIS K O C H B . S c , The University of Waterloo, Waterloo, 1992 A THESIS S U B M I T T E D I N P A R T I A L F U L F I L L M E N T OF T H E R E Q U I R E M E N T S F O R T H E D E G R E E OF D O C T O R OF P H I L O S O P H Y in T H E F A C U L T Y OF G R A D U A T E S T U D I E S (Department of Chemistry) We accept this thesis-as conforming to the^3^ed/standiB5i T H E U N I V E R S I T Y OF B R I T I S H C O L U M B I A October 1998 © I r i s Koch, 1998 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of CH6 fAl ^ T ^ V The University of British Columbia Vancouver, Canada Date QrA- S" -, W % DE-6 (2/88) ABSTRACT The determination of arsenic and antimony species in environmental samples can be used to assist in toxicity assessment, as well as to yield information about environmental processes. Such information about samples from the terrestrial environment was sought. Existing methods for speciation were adapted, including high-performance liquid chromatography ( H P L C ) coupled with inductively coupled plasma-mass spectrometry ( ICP-MS), for the determination of both arsenic and antimony species, and hydride generation-gas chromatography ( H G - G C ) with atomic absorption spectrometric ( A A S ) detection and mass spectrometric (MS) detection, for the determination of antimony species. Arsenate, when added to mycelial cultures of Scleroderma citrinum and Macrolepiotaprocera, was reduced to arsenite, but no further processes (i.e., methylation or formation of arsenosugars or arsenobetaine) were observed. This may indicate that the presence of more complex arsenicals in environmental mushroom specimens is dependent on symbiotic interactions between the fungus and its surroundings, rather than resulting from independent synthesis by the fungus. Pleurotus flabellatus oxidized antimony (III) to antimonate (Sb(OH) 6 ) , and formed an antimony-containing metabolite of unknown identity. Water soluble arsenic species were determined in a host of terrestrial and freshwater biota from a hot springs environment (Meager Creek, B C ) and from an area impacted by mining and smelting activities (Yellowknife, N W T ) . Arsenate and arsenite (the more toxic forms of arsenic) were the predominant species extracted from plants, mosses, microbial mats, algae and lichens. Small amounts of arsenosugars and methylated arsenic were detected as well. Arsenobetaine was discovered for the first time in lichens, and it was also the major form of arsenic in freshwater fish. The majority of detectable arsenic in freshwater mussels and snails was as arsenosugars and the tetramethylarsonium ion, respectively. Large amounts of arsenic, of an unknown ii toxicological and chemical nature, remained unextracted or undetected in all samples. A dimethylantimony compound was found in moss samples from Yellowknife, confirming that methylation of antimony takes place in the environment. TABLE OF CONTENTS Abstract i i Table o f Contents iv List o f Tables viii List of Figures x List of Abbreviations xiii Acknowledgment xv Chapter 1. I N T R O D U C T I O N 1 1.1. Flistorical aspects of arsenic and antimony 1 1.2. Chemistry of arsenic and antimony 2 1.3. Toxicity of arsenic and antimony compounds 6 1.4. Environmental chemistry of arsenic 7 1.4.1. Arsenic sources, uses and exposure to humans 7 1.4.2. Arsenic species in the marine environment 8 1.4.3. Arsenic species in the terrestrial environment 14 1.5. Environmental chemistry of antimony 16 1.5.1. Antimony sources, uses and exposure to humans 16 1.5.2. Antimony species in the environment 17 1.5.3. Biological and chemical transformations of antimony 18 1.6. Objectives and scope of work 20 References 22 Chapter 2. M E T H O D S F O R T H E A N A L Y S I S OF A R S E N I C A N D A N T I M O N Y 28 2.1. Methods for the Analysis of Arsenic 28 2.1.1. Introduction 28 2.1.2. Experimental 34 2.1.2.1. Chemicals and reagents 34 2.1.2.2. Instrumentation and methods of analysis 34 2.1.2.2.1. HPLC-ICP-MS 34 2.1.2.2.2. ESI-IT-MS 36 2.1.3. Results and Discussion 38 2.1.3.1. Speciation of arsenic compounds by H P L C 38 2.1.3.1.1. Anion exchange chromatography 38 iv r 2.1.3.1.2. Cation exchange chromatography 40 2.1.3.1.3. Ion-pairing chromatography 43 2.1.3.2. Characterization of arsenosugars and compounds in kelp powder extract 45 2.2. Methods for the Analysis of Antimony 65 2.2.1. Introduction 65 2.2.2. Experimental 67 2.2.2.1. Chemicals and reagents 67 2.2.2.2. Method of analysis for H G - G C - A A S 67 2.2.2.3. Sample preparation 70 2.2.3. Results and Discussion 71 2.2.3.1. Demethylation of trimethylantimony species in aqueous solution during analysis by using G C - H G - A A S 71 2.2.3.1.1. The effect of acid 71 2.2.3.1.2. The effect of concentration 74 2.2.3.1.3. The effect of sample matrix 75 2.2.3.1.4. Other studies to determine causes of demethylation 77 2.2.3.1.5. Suggested reasons for demethylation .78 References 81 Chapter 3. A R S E N I C A N D A N T I M O N Y I N M U S H R O O M S 86 3.1. Introduction 86 3.2. Experimental 88 3.2.1. Chemicals and reagents 88 3.2.2. Apparatus and method of analysis 90 3.2.2.1. HG-GC-AAS analysis for antimony speciation 90 3.2.2.2. HPLC-ICP-MS analysis for antimony and arsenic speciation 91 3.2.2.3. ICP-MS analysis for total arsenic and antimony concentrations 91 3.2.3. Cultivation of Pleurotusflabellatus fruiting bodies on a solid substrate 93 3.2.4. Preparation of pure submerged cultures of fungi 94 3.2.5. Sample preparation and analysis. 97 3.2.6. Isolation of an unknown compound containing antimony 98 3.3. Results and Discussion 100 3.3.1. Arsenic species in edible mushrooms 100 3.3.2. The interaction of arsenic species with pure submerged cultures of fungi 104 3.3.2.1. Culture experiments with Scleroderma citrinum 104 3.3.2.2. Culture experiments with Macrolepiota procera and Sparassis crispa 108 3.3.2.3. Summary of the interaction of arsenic with fungi that can produce mushrooms I l l 3.3.3. The interaction of antimony species with fungi 112 3.3.3.1. Cultivation of Pleurotus flabellatus fruiting bodies 112 3.3.3.2. Culture experiments with Pleurotus flabellatus 114 3.3.3.2.1. ICP-MS and HG-GC-AAS analysis of biomass extracts and media 114 3.3.3.2.2. HPLC-ICP-MS analysis of biomass extracts and media 117 3.3.3.3. Culture experiments with Scleroderma citrinum 126 3.3.3.4. Summary of the interaction of antimony with fungi that can produce mushrooms 129 References 131 Chapter 4. A R S E N I C I N T H E M E A G E R C R E E K H O T SPRINGS E N V I R O N M E N T 133 4.1. Introduction 133 4.2. Experimental 135 4.2.1. Chemicals and reagents 135 4.2.2. Sampling 135 4.2.3. Sample preparation and analysis 137 4.3. Results and Discussion 140 4.3.1. Total concentrations of arsenic in water samples 140 4.3.2. Total concentrations of arsenic in biota 141 4.3.3. Arsenic speciation in biota samples 145 4.3.3. 1. Algae and microbial mat samples 147 4.3.3.2. Vascular plants (sedge, cedar, fleabane, monkey flower) 152 4.3.3.3. Moss 155 4.3.3.4. Fungi including lichens 157 4.3.3.5. Extraction efficiency for arsenic species 158 4.3.4. Summary 161 References 162 Chapter 5. A R S E N I C I N T H E Y E L L O W K N I F H E N V I R O N M E N T 166 5.1. Introduction 166 5.2. Experimental 167 5.2.1 Chemicals and reagents 167 5.2.2. Sampling 168 5.2.3. Sample preparation and analysis 171 5.3. Results and Discussion 174 5.3.1. Water and sediment samples 175 5.3.2. Freshwater fish 176 5.3.2.1. Total arsenic in fish 177 5.3.2.2. Arsenic speciation in fish 179 5.3.3. Freshwater shellfi sh 187 5.3.3.1. Total arsenic in shellfish 187 5.3.3.2. Speciation of arsenic in shellfish 188 5.3.4. Plants 194 5.3.4.1. Total arsenic in plants 196 5.3.4.2. Arsenic speciation in plants 199 5.3.5. Algae and microbial mats 204 5.3.5.1. Total arsenic in algae 205 5.3.5.2. Arsenic speciation in algae 206 5.3.6. Mosses 209 vi 5.3.6.1. Total arsenic in mosses 209 5.3.6.2. Arsenic speciation in mosses 211 5.3.7. Lichens and mushrooms 214 5.3.7.1. Total arsenic in lichens and mushrooms 215 5.3.7.2. Arsenic speciation in lichens andfungi 217 5.4. Summary 229 References 231 Chapter 6. A N T I M O N Y I N E N V I R O N M E N T A L S A M P L E S 236 6.1. Introduction 236 6.2. Experimental 237 6.2.1. Chemicals and reagents 237 6.2.2. Sampling and sample preparation 237 6.2.3. I C P - M S analysis for total arsenic and antimony concentrations 241 6.2.4. H G - G C - A A S analysis for antimony speciation 241 6.2.5. H G - G C - M S analysis for confirmation of methylantimony species 242 6.3. Results and Discussion 244 6.3.1. Antimony species and total antimony in environmental samples 244 6.3.1.1. Inorganic antimony species 244 6.3.1.2. Methylated antimony species 247 6.3.1.3. Extraction efficiencies for biota and percent Sb species of total Sb in waters 250 6.3.1.4. Total concentrations of antimony compared with arsenic 251 6.3.2. The confirmation of antimony in samples containing methylantimony compounds by using H G - G C - A A S 253 6.3.3. The use of headspace H G - G C - M S for the speciation of antimony compounds 254 6.4. Summary 263 References 264 Chapter 7. C O N C L U S I O N S A N D F U T U R E W O R K 266 vii LIST OF TABLES Chapter 1 Table 1.1. Names, abbreviations and structures of some arsenic compounds 4 Table 1.2. Names, abbreviations and structures o f some antimony compounds 5 Chapter 2 Table 2.1. H P L C conditions for arsenic speciation 35 Table 2.2. Operation parameters for I C P - M S 36 Table 2.3. E S I - I T - M S experiments and fragments for pure arsenosugars 47 Table 2.4. E S I - I T - M S experiments for kelp powder extract 60 Table 2.5. Comparison of amounts of demethylation when using different concentrations of acids 74 Table 2.6. Studies to determine causes of demethylation 77 Chapter 3 Table 3.1. H P L C conditions for arsenic speciation 92 Table 3.2. H P L C conditions for antimony speciation 92 Table 3.3. Operation parameters for I C P - M S 93 Table 3.4. Summary of pure culture experiments 95 Table 3.5. Y M Broth ingredients and composition 96 Table 3.6. Arsenic species in edible mushrooms 101 Table 3.7. Comparison of proportions of arsenic species in mushrooms in the current study with those found in published studies 102 Table 3.8. Total concentrations of arsenic obtained by I C P - M S analysis for experiments conducted with S. citrinum 106 Table 3.9. Proportions of arsenic species (%) in experiments conducted with S. citrinum 107 Table 3.10. Arsenic species (% of arsenic extracted) found in wild specimens o f M procera and S. crispa (from Slejkovec etal.) 109 Table 3.11. Concentrations of arsenic species in experiments conducted w i t h M procera and S. crispa. 110 Table 3.12. Antimony in mushrooms after acid digestion, analyzed by hydride generation-GC-A A S 113 Table 3.13. Antimony extracted from mushrooms and soils (ppm dry weight), H G - G C - A A S analysis 114 Table 3.14. Antimony in media and biomass extracts of Pleurotus flabellatus grown in submerged culture 116 Table 3.15. Relative amounts of antimony compounds (%) in some P. flabellatus samples analyzed by using Method A 119 Table 3.16. Concentration of total antimony (ppm) in media and biomass extracts of Scleroderma citrinum grown in submerged culture 126 viii Chapter 4 Table 4.1. Operation parameters for I C P - M S 138 Table 4.2. H P L C conditions for arsenic speciation 139 Table 4.3. Some physical and chemical characteristics of Meager Creek waters 141 Table 4.4. Samples, sampling location, sampling times and arsenic levels in biota samples 142 Table 4.5. Estimated concentrations of arsenic species in biota samples 146 Table 4.6. Percent amounts of arsenic extracted 159 Chapter 5 Table 5.1. Operation parameters for I C P - M S 173 Table 5.2. H P L C conditions for arsenic speciation 173 Table 5.3. Concentrations of total arsenic and arsenic species in water samples and soil extracts 175 Table 5.4. Total arsenic concentrations in fish from location 9 178 Table 5.5. Comparison of arsenic concentrations by using protease and acid digestion methods for oyster tissue (NIST 1566) 180 Table 5.6. Concentrations of arsenic species in fish from location 9 181 Table 5.7. Moisture content and arsenic concentration in freshwater shellfish 188 Table 5.8. Concentrations of arsenic species in freshwater mussels and snails 189 Table 5.9a. Arsenic concentrations in plants from Yellowknife 196 Table 5.9b. Arsenic concentrations in plants from Yellowknife, previous study 197 Table 5.10. Concentrations of arsenic species in Yellowknife plants 200 Table 5.11. Percent arsenic species of total arsenic extracted from plants 201 Table 5.12. Arsenic concentrations in algae from Yellowknife 205 Table 5.13. Concentrations of arsenic species in Yellowknife algae 207 Table 5.14. Arsenic concentrations in mosses from Yellowknife 210 Table 5.15. Concentrations of arsenic species in Yellowknife mosses 212 Table 5.16. Arsenic concentrations in lichens and fungi from Yellowknife 216 Table 5.17a. Concentrations of arsenic species in Yellowknife lichens 218 Table 5.17b. Concentrations of arsenic species in Yellowknife fungi 219 Table 5.18. Proportions of total arsenic extracted, for lichens and mushrooms 222 Chapter 6 Table 6.1. Operation parameters for I C P - M S 241 Table 6.2. G C - M S parameters 243 Table 6.3a. Total antimony, extracted antimony species and estimated extraction efficiency in extracts of environmental biota samples 245 Table 6.3b. Total antimony, antimony species and percent Sb species of total Sb in environmental samples of water 246 Table 6.4. Comparison of total concentrations of antimony and arsenic for selected samples .252 Table 6.5. Relative amounts for methyantimony peaks in moss and water samples 254 ix LIST OF FIGURES Chapter 1 Figure 1.1a. The Challenger mechanism, showing alternating reducing and oxidative addition steps 9 Figure 1.1b. The structure for S-adenosylmethionine ( S A M ) 9 Figure 1.2. Pathway showing the formation of dimethylarsinoribosides and arsenobetaine 11 Figure 1.3. Pathway showing the formation of a trimethylarsonioriboside followed by the formation of arsenobetaine 13 Chapter 2 Figure 2.1. Chromatogram of standard arsenic compounds by using Hamilton PRP-X100 anion exchange column (15 cm) with 20 m M ammonium phosphate, p H 6 39 Figure 2.2. Chromatogram of standard arsenic compounds by using Whatman S C X cation exchange column with 20 m M pyridinium formate, p H 2.7 41 Figure 2.3. Chromatogram of standard arsenic compounds by using ion-pairing reversed phase chromatography, with G L Sciences C I 8 column and 10 m M T E A H / 4 . 5 m M malonic acid, p H 6.8, 0 . 1 % M e O H 44 Figure 2.4. Mass spectrum of arsenosugar X standard, M S - M S with m/z 329 selected, positive mode 49 Figure 2.5. Mass spectrum of arsenosugar X I standard, M S - M S with m/z 483 selected, positive mode '. 51 Figure 2.6. Mass spectrum of arsenosugar X I standard, M S - M S - M S with m/z 481 selected, then m/z 389, negative mode 52 Figure 2.7. Mass spectrum of arsenosugar XI I standard, M S - M S with m/z 393 selected, positive mode 55 Figure 2.8. Mass spectrum of arsenosugar X I I standard, M S - M S with m/z 391 selected, negative mode 56 Figure 2.9. Mass spectrum of arsenosugar XIII standard, M S - M S with m/z 407 selected, negative mode 57 Figure 2.10. Mass spectrum of arsenosugar XIII standard, M S - M S - M S with m/z 407 selected, then m/z 285, negative mode 58 Figure 2.11. Mass spectrum of precursor ion m/z 481 in kelp extract subjected to CID, negative mode 62 Figure 2.12a. Mass spectra for precursor ion m/z 407 in kelp extract subjected to CID, negative mode. Figure 2.12b. Precursor ion m/z 407 selected, then m/z 285 63 Figure 2.13. Schematic diagram for hydride generation of stibines, Method 3 69 Figure 2.14. Percent amounts of stibines generated from M e 3 S b C l 2 at varying pH, when using H G - G C - A A S 72 Figure 2.15a. Chromatogram of an aqueous fungus extract analyzed at neutral p H (water only, unbuffered). Figure 2.15b. Chromatogram of the same aqueous fungus extract analyzed at p H 6.2, citrate buffer. Figure 2.15c. Chromatogram of antimony-free aqueous fungus extract spiked with 200 ng M e 3 S b C l 2 , analyzed as in Figure 2.15a. Figure 2.15d. Chromatogram of 200 ng M e 3 S b C l 2 in water analyzed as in Figure 2.15a 76 Chapter 3 Figure 3.1. Chromatograms of standard antimony compounds (100 ppb each) on two H P L C -I C P - M S systems 118 Figure 3.2. Chromatogram (Method A ) of medium after 14 days of growth for P. flabellatus amended with Sb (V) 120 Figure 3.3. Chromatograms of P. flabellatus media and biomass extracts (Method B ) for experiments amended with Sb (V) 122 Figure 3.4. Chromatograms of unknown B 4 (Method B 124 Figure 3.5. Raw area counts for Sb(OH)6~ in media for P. flabellatus grown in Sb (Ill)-amended culture 125 Figure 3.6. Chromatograms for biomass extracts (fresh weight) for S. citrinum experiments (Method A ) 128 Chapter 4 Figure 4.1. Map (not to scale) of Meager Creek Hot Springs area showing sampling locations 136 Figure 4.2. Chromatograms for Algae 1 and laboratory standards showing the presence of arsenosugars X and X I 148 Figure 4.3. Chromatograms of a microbial mat extract (top layer, microbial mat) and extract spiked with arsenosugar X I 149 Figure 4.4. Seasonal arsenic speciation in higher plants, sedge (Scirpus sp.) and fleabane (Erigeron sp.) 154 Figure 4.5. Seasonal and spatial arsenic speciation in moss (Fumaria hygrometrica) 157 Chapter 5 Figure 5.1a. Map of Royal Oak Giant mine property and surrounding area 169 Figure 5.1b. Map of Yellowknife, showing Royal Oak Giant Mine and Miramar Con Mine.. . . 170 Figure 5.2. Chromatogram of protease digest (PD) of a Yellowknife fish 184 Figure 5.3. Relative amounts of arsenic species in moss and associated organisms from Yellowknife ( Y K ) and Meager Creek ( M C ) 213 Figure 5.4. Arsenobetaine ( A B ) in lichens 220 Figure 5.5. Relative amounts of arsenic species in the puffball mushroom Lycoperdon sp. from location 7 (Giant Mine tailings pond) and location 15 (Con Mine tailings pond) 223 xi Figure 5.6. Chromatogram ofPaxillus involutus extract (diluted lOx), showing the presence of unknown compounds 225 Chapter 6 Figure 6.1a. Map (not to scale) of Meager Creek Hot Springs area showing sampling locations 239 Figure 6.1b. Map of Yellowknife sampling locations 240 Figure 6.2. Chromatogram obtained by using H G - G C - A A S (217.6 nm) showing stibines generated at neutral p H from 100 m L of a sample of standing water from location 4 in Yellowknife 248 Figure 6.3. Chromatogram mass spectrum following H G - G C - M S for M e 3 S b generated from 30 n g M e 3 S b C l 2 255 Figure 6.4. Chromatogram mass spectrum following H G - G C - M S of stibines generated from 100 ng M e 3 S b C l 2 (1 M HC1) 257 Figure 6.5. Chromatogram mass spectrum following H G - G C - M S of moss extract (June, Y K Location 1) ..259 Figure 6.6. Chromatogram mass spectrum following H G - G C - M S of moss extract (August, Y K Location 1) 260 Figure 6.7. Chromatogram and mass spectra, following H G - G C - M S of a moss extract (August, Y K Location 4) 261 Figure 6.8. Chromatogram and mass spectra, following H G - G C - M S of a snail extract 262 xii L I S T O F A B B R E V I A T I O N S A B arsenobetaine A C arsenocholine AsS extract containing arseno sugars B C F bioconcentration factor B T T Bi l l ' s Toxic Team C E capillary electrophoresis C E P A Canadian Environmental Protection Agency CID collision induced dissociation C R Campbell River C R M certified reference material D a Dalton (atomic mass unit) D E digestion efficiency D M A dimethylarsinate D M A A dimethylarsinic acid D M A E dimethylarsinylethanol D S A dimethylstibinic acid dw dry weight E D T A ethylenediaminetetraacetic acid E E extraction efficiency ESI - IT-MS electrospray ionization-ion trap-mass spectrometry F A A S flame atomic absorption spectrometry F A B fast atom bombardment FID flame ionization detection fw fresh weight G F A A S graphite furnace atomic absorption spectrometry H G - G C - A A S hydride generation-gas chromatography-atomic absorption spectrometry H P L C high performance liquid chromatography i.d. inner diameter I C P - M S inductively coupled plasma-mass spectrometry IS ionspray L O D limit o f detection M + H molecular ion + H M - H molecular ion - H M / W methanol/water (1:1) extraction m/z mass to charge ratio M C Meager Creek M e O H methanol M M A methylarsonate M M A A methylarsonic acid M S mass spectrometry M S A methylstibonic acid M S " tandem mass spectrometry N I S T National Institute for Science and Technology o.d. outer diameter xiii O E S optical emission spectrometry P C pixie cups P D protease digestion ppb parts per billion, ng/g, pg/kg, or ug/L ppm parts per million, ug/g, mg/kg, or mg/L P T F E poly(tetrafluoroethylene) R ratio obtained from (fresh weight mass)/(dry weight mass) R B F round bottom flask S A M S-adenosylmethionine SD standard deviation SFC supercritical fluid chromatography SUDS Sudden Infant Death Syndrome T E A H tetraethylammonium hydroxide T M A trimethylarsenic (unspecified structure) T M A O trimethylarsenic oxide T R A time resolved analysis TSbO trimethylantimony oxide U K unknown compound U S - E P A United States Environmental Protection Agency U S - F D A United States Food and Drug Administration U V ultra violet v/v volume per volume w/v weight per volume W H O World Health Organization X - X I I I arsenosugars X through XIII Y K Yellowknife xiv A C K N O W L E D G M E N T I wil l never be able to express how deeply grateful and warmly attached I feel towards all the people who have been a part of my life for the last five years. M y life would not have been enjoyable or productive without them. First, I would like to thank my supervisor, B i l l Cullen, who kept me inspired when I most needed it. I am also very grateful to my supervisory committee, including Mike Blades, Guenther Eigendorf, Tom Pederson, and especially David Chen. M y second supervisor, Ken Reimer, gave me many opportunities to travel and expand my skills, and I thank him for all his help. Other people at R M C have been invaluable to me, including Chris Ollson, Ian Mace, Chris Knowlton, Wayne Ingham, Dave Pier and many of my other friends there. The people in Yellowknife without whom the Yellowknife sampling trips would not have happened are Steve Harbicht at Environment Canada and Steve Schultz from Giant Mine, among others. I have learned much about fields other than chemistry and I am very grateful to Elena Polishchuk and Paul Haden for their expertise and excellent teaching skills. I am also indebted to the experts who helped me with identification of biological samples, including Dr. W . B . Schofield, Olivia Lee, Julie Oliveira, James Black and Mike Fournier. I would also like to thank Martin Tanner and Paul Morgan for the use of the French press. I am particularly grateful to Bert Mueller and his expertise with the I C P - M S . I am indebted to the present and former members of my research group, for many fruitful discussions and collaborations, as well as friendship. They include Bianca Kuipers, Vivian Lai , Corinne Lehr, Paul Andrewes, L ix ia Wang, Changqing Wang, Kirsten Falk, Dietmar Glindemann, Sepp Lintschinger, Jorg Feldmann, Andrew Mos i , Chris Harrington, Spiros Serves, and especially Chris Simpson. Summer students who have been a great help are Meghan Winters and L i n Tran. Some of the above people were kind enough to read through parts of my thesis and I thank them, as well as Graham Cairns, for it. Finally I would like to thank my friends, my sister and my parents for their love and support, and without whom I would not have reached my goal. XV Chapter 1 INTRODUCTION "Any wild place is filled with incredible things happening." David Cavagnaro, This Living Earth The chemistry of the environment is a complex area of study, reaching into the fields of biology, geology, physics, toxicology and medicine, among others. Two crucial aspects of environmental chemistry are the identification of chemical contaminants and the determination of their fate in natural and anthropogenic environments. This knowledge is often used to discover and solve problems caused by the presence of chemical contaminants. Such problems become of utmost concern when they include adverse health effects in humans, animals or plants. Metals or metalloids are often found in the environment as chemical contaminants. Two examples are arsenic and antimony, which are closely related in chemical behaviour. 1.1. Historical aspects of arsenic and antimony Both arsenic and antimony have had paradoxical uses throughout history, as poisons and as panaceas. Arsenic trioxide, known as "white arsenic" is historically one of the most common poisons. Strangely, this same compound was used until recently in a region in Austria by the "arsenic eaters", who attempted to increase their strength, virility and longevity by ingesting arsenic trioxide daily 1. Hippocrates (460-377 B .C . ) recommended the use of a paste of realgar (As 4 S 4 ) to treat ulcers 2. In the early part of this century (1900-1950), organoarsenic compounds, such as salvarsan, neosalvarsan and atoxyl, were used for the treatment of syphilis and sleeping sickness2. Lewisite ( C l C H = C H A s C l 2 ) , a severe blistering agent, was developed for chemical warfare purposes. Arsenic trioxide is thought to be the active ingredient in current traditional Chinese treatments of arthritis, skin disorders and even cancer3. Antimony also has a fascinating history. It is mentioned in the Bible as a cosmetic and it was commonly used up until the mid-18 t h century as a medicine to produce sweating, as an emetic, as a purgative, or all three4. Many preparations of antimony are described in the well-known "The Triumphal Chariot of Antimony" by Basil Valentine, first published in 16045. Fatalities resulting from its use did not prevent its mention in Martindale's Extra Pharmacopoeia, as recently as 1941 4 Antimony was of great interest to alchemists, and one of its properties was that it "united with or devoured all the metals then known, with the exception of gold" 6 . Some cases of alleged deliberate antimony poisoning also have been documented4. 1.2. Chemistry of arsenic and antimony Arsenic and antimony are found in Group 15 of the periodic table. A s metalloids, they possess characteristics of both metals and non-metals. Antimony is more metallic than arsenic because it can exist as aqueous cationic species, [SbO +] and [Sb0 2 + ] (but not free [Sb 3 +] or [Sb 5 +]), and its trioxide is amphoteric, dissolving in both strong acid and strong base. Arsenic, on the other hand, forms oxy-anionic species only. Its trioxide is acidic and hence dissolves in base, which is characteristic of non-metals7. Both arsenic and antimony form hydrides, implying that, like non-metals, they can also exist in a negative oxidation state. 2 Arsenic and antimony possess five valence electrons like other group 15 elements. Their four oxidation states are -3, 0, +3 and +5. A n antimony compound with composition SD2O4 has been observed, implying that a +4 oxidation state is possible, but this oxide has been shown to contain only S b v and Sb f f l atoms8. The arsenic and antimony compounds that are pertinent to this thesis are either hydrides, organometallic compounds (possessing a A s - C or Sb-C bond), oxyanions, or complexes of the inorganic cations. Table 1.1 and Table 1.2 show names, abbreviations and structures for some arsenic (Table 1.1) and antimony (Table 1.2) compounds. 3 Table 1.1. Names, abbreviations and structures of some arsenic compounds. Name Abbreviation Structure/Formula Arsenic acid, arsenatea A s ( V ) As(0)(OH) 3 Arsenous acid, arsenite3 As (III) As (OH) 3 Monomethylarsonic acid" M M A C H 3 A s O ( O H ) 2 Dimethylarsinic acid a D M A ( C H 3 ) 2 A s O ( O H ) Trimethylarsine oxide b T M A O ( C H 3 ) 3 A s O Methylarsine M e A s H 2 C H 3 A s H 2 Dimethylarsine M e 2 A s H ( C H 3 ) 2 A s H Trimethylarsine M e 3 A s ( C H 3 ) 3 A s Tetramethylarsonium ion b M e 4 A s + ( C H 3 ) 4 A s + Arsenobetaineb A B ( C H 3 ) 3 A s + C H 2 C O O " Arsenocholine b A C ( C H 3 ) 3 A s + C H 2 C H 2 O H Dimethylarsinylethanol D M A E ( C H 3 ) 2 A s ( 0 ) C H 2 C H 2 O H Arsenosugars c X - X I I I See figure below o I  ( C H 3 ) 2 A s C H 2 OH I O C H 2 C H C H 2 — R Sugar X X I X I I XIII R - O H - O P 0 3 H C H 2 C H ( O H ) C H 2 O H -S0 3 H -OS0 3 H a These compounds are commercially available. b These compounds are available in our lab, having been synthesized according to common methods (see Chapter 2). c These compounds are available in our lab; they are found in a laboratory reference material (kelp powder) and have also been donated (see Chapter 2). d Numbering system (X-XIII) according to Shibata et al9 Table 1.2. Names, abbreviations and structures of some antimony compounds. Name Antimonate 3 Pentavalent inorganic antimony Antimony trioxide" Antimony trisulfide Antimony potassium tartrate3 Trivalent inorganic antimony Methylstibonic acid Dimethylstibinic acid Trimethylantimony oxide Trimethylantimony dihydroxide b Trimethylantimony dichloride b Methylstibine Dimethylstibine Trimethylstibine Abbreviation S b ( V ) Sb (V) Sb (III) Sb (III) Sb (III) M S A D S A TSbO M e 3 S b ( O H ) 2 M e 3 S b C l 2 M e S b H 2 M e 2 S b H Me 3 Sb Structure/Formula Sb(OH)<f inorganic v S b (complexed) Sb 2 0 3 , Sb(OH) 3 ( a q ) Sb 2 S 3 K2 \ / \ / C H - O O - C H C H - O O - C H / w_ \ 0 ^C - 0 - S b - 0 ^ C ^ 0 inorganic m S b (complexed) C H 3 S b O ( O H ) 2 (CH 3 ) 2 SbO(OH) (CH 3 ) 3 SbO (CH 3 ) 3 Sb(OH) 2 ( C H 3 ) 3 S b C l 2 C H 3 S b H 2 ( C H 3 ) 2 S b H (CH 3 ) 3 Sb a These compounds are commercially available. b These compounds are available in our lab, having been synthesized according to common methods (see Chapter 2). 5 1.3. Toxicity of arsenic and antimony compounds Although the word "arsenic" is usually immediately associated with poison, the actual toxicity of arsenic in a sample is dependent on the chemical form that it takes. The same principle applies for antimony. Arsenic and antimony compounds in the -3 oxidation state are more toxic than those in the +3 oxidation state, which are more toxic than those in the +5 oxidation state. Organometallic compounds in the +5 oxidation state are less toxic than inorganic ones and some compounds, like arsenosugars and arsenobetaine, have not exhibited any toxic behaviour at all in the systems tested 1 0 ' 1 1 ' 1 2. The dependence of toxicity on the chemical form (or species) of arsenic and antimony makes their identification (i.e., speciation analysis) necessary. Trivalent arsenic is thought to exert its toxicological effects by binding with sulfhydryl groups on enzymes 2 ' 1 3. Inhibition of pyruvate dehydrogenase takes place for both trivalent arsenic and trivalent antimony 1 3 ' 1 4. Pentavalent arsenic is thought to compete with phosphate during phosphorylation to form unstable arsenyl esters, interfering with bioenergetic processes 2' 1 3. Trivalent antimony binds easily in vivo to sulfhydryl groups, such as those on enzymes, most likely constituting the toxic action of these compounds 1 5 ' 1 4. Pentavalent antimony is excreted rapidly from the body but it has been shown that the liver can reduce Sb (V) to Sb (III) 1 6. The International Agency for Research on Cancer ( IARC) has determined that there is sufficient evidence to consider arsenic a human carcinogen, and to consider antimony trioxide an animal (but not human) carcinogen 1 7. 6 1.4. Environmental chemistry of arsenic 1.4.1. Arsenic sources, uses and exposure to humans Arsenic is the twentieth most abundant element in the Earth's crust, and is often associated with sulfidic ores such as arsenopyrite (FeAsS), enargite (CU3ASS4), orpiment (As 2 S 3 ) , realgar (AS4S4) and proustite ( A g 3 A s S 3 ) 1 3 ' 1 8 . Arsenic is usually recovered as arsenic trioxide from the processing of ores, and its major present uses are in the production of arsenic-containing agricultural pesticides (including disodium methylarsonate, sodium methylarsonate, methylarsonic acid ( M M A ) and dimethylarsinic acid ( D M A ) ) ; wood preservatives (including chromated copper arsenate ( C C A ) , ammoniacal copper arsenate ( A C A ) and fluorchrome arsenate phenol (FCAP)) ; and animal feed additives (including arsanilic acid and 4-nitrophenylarsonic acid) 1 3 ' 1 9 . Arsenic is also used as a doping agent in solid-state products 1 9. Arsenic can enter the environment anthropogenically as a consequence of its industrial use, from mining and smelting operations and through the application of arsenic-containing pesticides. Natural inputs of arsenic to the environment can result from the weathering of rocks and geothermal activities, leading to open ocean arsenic concentrations typically ranging from 18 18 70 0.0056-11 ppb and terrestrial soil and rock concentrations ranging from 0.4-100ppm ' . Humans can be exposed to arsenic through inhalation, especially for workers in industries utilizing arsenic, through food (such as fish, shellfish, or marine algae) and through drinking water (e.g., in India, groundwater supplies contain elevated levels of arsenic 2 1). Arsenic is considered to be a priority pollutant by the United States and the Canadian Environmental Protection Agencies ( U S - E P A and C E P A) . U S - E P A and C E P A permit drinking water levels to be a maximum of 50 2 2 and 25 ppb 2 3 (u,g/L), respectively. The World Health 7 Organization (WHO) recommends a daily limit through food of 0.05 mg total arsenic/kg body weight 2 2, but for inorganic arsenic a weekly limit of only 15 ug As/kg body weight is suggested' 1.4.2. Arsenic species in the marine environment The major species of arsenic in seawater is As ( V ) 2 5 ' 2 6 , although sometimes as much as 10-30% of the total arsenic takes the form of As (III), M M A and D M A in waters from the euphotic zone, in deeper ocean waters and in interstitial waters 2 0 ' 2 5 ' 2 6 . Arsenate is thermodynamically predicted to be the major compound in oxygenated seawater (pH 8) 2 0 and therefore the presence of As (III) and methylated species imply that biotransformation is taking place. This is indeed observed to be the case because marine algae 2 7, bacteria and fungi 2 0 methylate arsenic. The Challenger mechanism 2 8 (Figure 1.1a), involving alternating steps of reduction and oxidative addition of a methyl group, is considered to be a possible methylation pathway. Studies have shown that S-adenosylmethionine ( S A M ) (Figure l i b ) acts as a methyl donor 2 0. 8 1.1a AsO(OH) 3 2 6 » As (OH) 3 C h 3 » C H 3 A s O ( O H ) 2 C H 3 A s ( O H ) 2 CH3As(OH) 2 ^ (CH 3)2AsO(QH) 2 e ~ » (CH 3 ) 2 As(OH) V (CH 3 ) 3 AsO (CH 3 ) 3 AsO 2 e " » (CH 3 ) 3 As O H OH Figure 1.1a. The Challenger mechanism , showing alternating reducing and oxidative addition steps. Figure 1.1b. The structure for S-adenosylmethionine ( S A M ) ; the circled methyl group is donated during the methylation steps in the mechanism. In marine sediments arsenic concentrations can range from 0.4-455 ppm 1 8 . The majority of arsenic in sediment is associated with Fe and M n oxides and some is associated with carbonates and organic material 2 0. M M A and D M A have been found in environmental marine sediment samples2 0. Laboratory experiments conducted aerobically and anaerobically with cultures from marine sediments, amended with M M A and D M A , suggest that tetramethylarsonium ion may be formed and thus may be present in trace quantities2 9. Reliable evidence for the presence of arsenobetaine (but not A C , T M A O or M e 4 A s + ) in environmental 'if) estuanne waters now exists . 9 Marine algae contain levels of arsenic that are considerably higher than the levels in the surrounding water, ranging from 0.8 to 12.1 ppm wet weight 3 1. The major water-soluble forms of arsenic in most species of marine algae are the arsenosugars, some of which are shown in Table l . l 3 1 ' 3 2 . One exception is a Japanese edible algae, hijiki (Hizikiafusiforme), which contains 50% of its arsenic as arsenate33. Marine animals also contain higher levels of arsenic compared to the levels in the surrounding water 3 1, although biotransformation studies via the marine food chain indicate that biomagnification of arsenic probably does not take place 3 4. In contrast to most previous studies, one study has shown that biomagnification takes place in a short marine food chain (seaweed -> herbivorous marine gastropod - » carnivorous marine gastropod) 3 5. Arsenobetaine is the major arsenic compound in most marine animals, although the tetramethylarsonium ion has been found in marine snails and clams, arsenocholine has been found in gastropods and dogfish muscle 3 6, trimethylarsine oxide has been found in fish, and arsenosugars have been found in bivalve mollusks 3 1. A pathway for the formation of arsenosugars, and, from them, arsenobetaine, has been proposed and is shown in Figure 1.23 2'3 7. 10 (CH 3)2AsO(OH) 2 e ~ > (CH 3 ) 2 As(OH) SAM N HoN (CH 3 ) 2 As(OH) + C H 3 + S - h C H 2 . 0 , < OH OH NH, O ( C H 3 ) 2 A s C H 2 v N . 0 N ^ N M ( 2 - 1 ) OH OH O ( C H 3 ) 2 A s C H 2 . /O/QR O R + ( C H 3 ) 2 A s C H 2 . . 0 N OH OH OH O (CH 3 ) 2 As . [O] (2.2) X>H 2e~, then I C H 3 + O (CH 3 ) 2 As . X O O " (2.3) 2e", then C H 3+ (CH 3 ) 3 As-*0H Arsenocholine (AC) [O] ( C H 3 ) 3 A s v ^ ^ C O O " Arsenobetaine (AB) Figure 1.2. Pathway showing the formation of dimethylarsinoribosides and arsenobetaine30. The pathway for the synthesis of arsenosugars is similar to the Challenger mechanism up until the formation of D M A (the starting molecule in Figure 1.2). After the reduction of D M A to ( C H 3 ) 2 A s O H , S-adenosylmethionine ( S A M ) is proposed to be the donor of the ribosyl moiety. The intermediate 2.1 in Figure 1.2 has been isolated from the kidney of the clam Tridacna maxima (symbiotic algae grow in the mantle of the clam and algal products accumulate in the kidney), providing support for this pathway 3 8. The formation of arsenobetaine from arsenosugars, as proposed in Figure 1.2, involves an intermediate compound 2.2 (dimethylarsinoylethanol, D M A E ) , which has not yet been identified as a naturally occurring compound 3 2. However, trace amounts of dimethylarsinoylacetic acid (2.3), as well as arsenosugars and arsenobetaine, have been reported in a mussel sample 3 9, which may lend support to one of the routes in the proposed pathway in Figure 1.2. Small amounts of a trimethylarsonioriboside (Figure 1.3, 3.1) have been identified in algae 4 0 leading to the proposal of an alternate pathway for the formation of arsenobetaine, shown in Figure 3. The formation of arsenobetaine from the trimethylarsonioriboside has been demonstrated to be a facile process in laboratory experiments 4 1 ' 4 2, which may indicate that although the levels of the starting trimethylarsonioriboside are very low in algae, this pathway may, to some extent, contribute to the formation of arsenobetaine in the marine environment3 2. 12 o (CH 3 ) 2 AsCrW ,0 OCH2CH(OH)CH 2OS0 3~ OH OH 2e" ( C H 3 ) 2 A s C H 2 \ .0 .OCH 2CH(OH)CH 2OS0 3" OH OH CH 3+ ( C H 3 ) 3 A s+ C H 2 \ / 0 'VOCH 2CH(OH)CH 2 OS0 3 " OH OH (3-1) ( C H 3 ) 3 A s V / \ J C ^ On (CH 3 ) 3 AsV v / COO" Arsenocholine (AC) Arsenobetaine (AB) Figure 1.3. Pathway showing the formation of a trimethylarsonioriboside (3.1) followed by the formation of arsenobetaine30. 13 M u c h is known about arsenic species in the marine environment, and reviews by Cullen et al.20 and Francesconi et al.32 address this topic in more detail than that given here. Such a wealth of knowledge is available because of the ubiquitous presence of arsenic in marine samples, as well as sufficiently high levels of arsenic for reliable speciation analysis. However, the information that is obtained is dependent on the analytical methods available and as a result, little reliable data exists for arsenic compounds that are not water soluble. Additionally, the presence of trace amounts of crucial intermediates in the formation of organoarsenicals in environmental samples is dependent on the detection limits possible by using current analytical methods. 1.4.3. Arsenic species in the terrestrial environment Arsenic concentrations in freshwater systems vary depending on the geological composition of the area and the input from anthropogenic sources, such as agriculture or mining operations. Levels in rivers and lakes range from 0.1 to 75 ppb in some reports 4 3, and they can be substantially elevated in hot springs (up to ppm levels) 4 4. A s (V) is usually the major species, but high proportions (30-75% of total arsenic) of As (III) have been known to occur 2 1 ' 4 5 , as well as M M A , D M A , T M A O and trivalent methyl- and dimethylarsenic species 4 6 ' 4 7. Methylation of inorganic arsenic by anaerobic microorganisms associated with lake sediments has been observed 4 8, and biomethylation is expected to take place following the pathway proposed by Challenger (Figure 1.1). In soils, arsenic is associated with Fe and A l compounds 4 9. The amount of arsenic in the fraction that is soluble and hence available to plants has been estimated to be 0.07-0.2% of the estimated total, and the species of arsenic in this fraction were found to be As (V) and As (III) 5 0. Another study showed that from 9 soil samples, only one sample contained arsenite in a proportion greater than 1.38% of the estimated total arsenic, and As (V) was assumed to constitute the remainder5 1. However, arsenic in terrestrial soil and sediment is not limited to inorganic forms only; ( C H 3 ) A s H 2 , ( C H 3 ) 2 A s H , ( C H 3 ) 3 A s , ( C 2 H 5 ) ( C H 3 ) 2 A s , ( C 2 H 5 ) 2 ( C H 3 ) A s and ( C 2 H 5 ) 3 A s were generated with sodium borohydride from river sediments5 2. As wil l be discussed in Chapters 4 and 5, few studies have examined the speciation of arsenic compounds in plants and other terrestrial biota, and details of previous studies are given in sections 4.3 and 5.3. A Japanese study concludes that methyl species, of unknown chemical structure, are present in a green alga, a diatom, a freshwater prawn, a marsh snail, freshwater fish and fly larvae, sampled from an area impacted by geothermal waters and containing elevated arsenic levels 5 3. Mostly inorganic species, some D M A and trace amounts of M M A and arsenobetaine are present in ants living in an arsenic contaminated area 5 4. Vegetables grown in arsenic amended soil were found to contain mostly arsenate, but trace amounts of M M A were present in lettuce, potato and swiss chard 5 5. The findings of arsenobetaine and arsenocholine, but not arsenosugars, in mushrooms 5 6 ' 5 7 ' 5 8 ' 5 9 led to the postulation by some researchers that arsenosugars are not involved in the biosynthesis of arsenobetaine32. Since the discovery of arsenosugars in the terrestrial environment 6 0 ' 6 1, however, the possibility has emerged that pathways (e.g., Figure 1.2) in the terrestrial environment are similar to those in the marine environment. Most studies to date indicate that arsenobetaine and arsenosugars are not as common in the terrestrial environment as they are in the marine environment. Among mushrooms, for example, the form of arsenic varies greatly from one species to the next. Some species contain only inorganic arsenic, and others contain only arsenobetaine. Clearly, studies to determine the arsenic compounds present in terrestrial samples are necessary to elucidate the chemical processes taking place in the terrestrial environment. 15 1.5. Environmental chemistry of antimony 1.5.1. Antimony sources, uses and exposure to humans Antimony occurs at about one-tenth the concentration of arsenic in the Earth's crust, and is usually found as stibnite (Sb 2 S 3 ) , as well as in ores of copper, silver and lead 6 2 . The major use of antimony is in the form of Sb 203, as a flame retardant in textiles, paper and plastics 6 2 ' 6 3. Antimony compounds are used to a lesser extent in paints and ceramics; as catalysts, glass decolourizers and metal hardeners; and in the semiconductor industry 6 2 ' 6 4. Antimony compounds are also used as a treatment for tropical parasitic diseases 6 5' 6 6. Sources of antimony in the environment are usually similar to those for arsenic, since antimony often occurs together with arsenic-containing ores 6 7. Hence, weathering of rocks in areas with high arsenic and antimony content, geothermal activities, mining and smelting operations, and industries utilizing antimony can all contribute to the introduction of antimony into the natural environment. For example, high levels of antimony relative to other metalloids have been observed in landfill and sewage sludge fermentation gases 6 8 ' 6 9. Humans are not frequently exposed to antimony, except in working conditions that involve antimony, such as battery charging 6 4, antimony processing, and welding 6 6 . Usually, ecological exposure to antimony is also accompanied by exposure to other toxic compounds, such as those of lead, arsenic, cobalt or silica 6 6 . Nevertheless, the U S - E P A considers antimony to be a priority pollutant 7 0 ' 7 1 and the threshold limit permitted for antimony and its compounds in work room air is 0.5 mg Sb/m 3 7 0 . The United States Food and Drug Administration (US-FDA) tolerates a maximum of 2 ppm of antimony in food 6 6 , and the accepted daily limits for humans 16 orally exposed to antimony compounds (over an extended period of time) range from 24.5-32.5 u.g of antimony compound per day 6 2. 1.5.2. Antimony species in the environment Sb (V) species are thermodynamically most favourable under aerobic conditions and hence Sb(OH)6~ is predicted to be the predominant species of inorganic antimony in oxygenated water 7 2. Sb (III) compounds are thermodynamically predicted to be oxidized to Sb (V) at neutral p H and in an oxidizing environment. Sb 203 dissolves to a limited extent in water at neutral p H (<5 ppm), forming an undissociated species, that is, H S b 0 2 or Sb(OFf)37 2. The structure of organoantimony compounds in aqueous solution is unknown, except for M e 3 S b C l 2 , M e 3 S b B r 2 and Me 3 SbI 2 , which hydrolyze at neutral p H to form Me 3 Sb(OH) 2 , and possibly M e 3 S b 0 7 3 ' 7 4 . In accordance with the thermodynamic prediction, most studies report predominantly Sb (V) with small proportions of Sb (III), methyl- or dimethylantimony species in seawater 7 5 ' 7 6, estuarine waters 7 5 ' 7 7 , r ivers 7 5 ' 7 8 ' 7 9 , waste waters 8 0 ' 8 1, geothermal waters 8 2 and condensed water from landfill gas 6 9. Levels of antimony in waters are typically less than 1 ppb 6 2 , although polluted waters can contain levels of antimony from 300-800 ppb 7 5 ' 8 1 . Sb (V) as Sb(OH) 6" was the major species extracted from soils 8 2, and it was postulated that M e 3 S b O was present in one such extract 8 0. Compounds forming C H 3 S b H 2 , ( C H 3 ) 2 S b H , (CH 3 ) 3 Sb and ( C 2 H 5 ) 3 S b were found in river sediment that was reacted with sodium borohydride 5 2, indicating the presence of small, but detectable levels of organoantimony compounds in soils and sediments. Antimony was shown to be associated with iron and aluminum in sediments8 3, but in soils it was mostly in a form not extractable by sequential leaching procedures; the highest extractable amount was bound to Fe -Mn oxides 8 4. Antimony 17 concentrations of up to 1489 ppm have been found in soil samples near an antimony smelter . In contrast, the levels of antimony in the Earth's crust are estimated to be between 0.2 and 1 ppm 6 6 . Less than 1% of antimony can be extracted from soil by using an aqueous extraction method 8 2, so it is not surprising that levels in human fluids were not significantly elevated, with respect to controls, following internal exposure to antimony contaminated soi l 8 6 . The volatile antimony compounds S b H 3 and MesSb have been found in gases sampled above hot springs, landfills and from sewage 6 8 ' 6 9 ' 8 7. Very few studies have speciated antimony in biota. Dodd et al. found methylated antimony species in aquatic plants collected from an area impacted by mining and hence containing elevated levels of antimony 8 8. Kantin reported mostly Sb (V) in extracts of marine algae, with low levels of Sb (III) in some samples8 9, and similar results were found in extracts of mollusk shells, with only one sample containing Sb (III) 9 0. Total levels of antimony were found to be elevated in biota collected near an antimony smelter with respect to those collected from control areas, and the authors concluded that uptake, but not biomagnification, of antimony takes place 8 5 ' 9 1 . 1.5.3. Biological and chemical transformations of antimony The findings of methylated antimony compounds in environmental samples, as mentioned in section 1.5.2, suggest that methylation is taking place in the environment. Recently, it was confirmed that biological methylation of antimony takes place by anaerobic soil cultures 9 2 and, to a lesser extent, by aerobic cultures of the fungus Scopulariopsis brevicaulis93,94. The volatile antimony compound formed, Me3Sb, was found to oxidize rapidly in an aerobic environment9 4; soluble dimethyl- and trimethylantimony compounds were found in liquid culture media 9 3. S. brevicaulis was capable of converting only 0.001-0.01% of antimony in liquid culture to 18 Me 3 Sb , which is much less than its ability to convert arsenic to M e 3 A s (~ 1%) . The volatilization of antimony by S. brevicaulis from mattresses contaminated with the fungus and containing antimony as a flame retardant has been suggested to be a cause of the sudden infant death syndrome (SIDS), or cot death 9 5 ' 9 6. However, stibine, postulated to be the volatile compound responsible for the antimony poisoning, has not yet been found as an antimony metabolite formed by S. brevicaulis cultures amended with antimony. This observation, together with the low conversion of antimony to volatile compounds by the fungus, makes it unlikely that antimony is linked to SIDS. Additionally, reliable epidemiological evidence linking the presence of antimony in mattresses to victims of SIDS is lacking 9 7 . Sb (III) is oxidized to Sb (V) by fungal cultures 9 8 as well as by the freshwater alga Chlorella vulgaris". The antimony that was accumulated by C. vulgaris cells appears to be bound to proteins whose molecular weight is around 4 x 10 4 D a 9 9 . The bacterium Stibiobacter senarmontii is reported to use S b 2 0 3 as a substrate for growth and may also convert this compound to Sb (V). Another bacterium Thiobacillus ferrooxidans has been observed to oxidize stibnite 1 0 0. Trimethylstibine is insoluble in water, but it quickly and spontaneously forms the water soluble compound M e 3 S b O in the presence of oxygen. This is postulated to contribute to the mobilization of M e 3 S b in the environment 1 0 1. Trimethylstibine was observed to form the water soluble [Me4Sb+] ion in the presence of alkyl halides (which may be present in the environment) in polar solvents, leading to the speculation that this may also be a process occurring in the aqueous environment 1 0 2. N o evidence exists yet for the presence of the [Me 4 Sb + ] ion in the environment. The lack of analytical methodology for the trace detection of such antimony species likely limits the range of compounds found in the environment. 19 1.6. Objectives and scope of work The general theme in this work is to expand the knowledge about the species of arsenic and antimony occurring in the terrestrial environment. The goal of Chapter 2 is to describe improvements and clarifications of some existing analytical methods for arsenic and antimony compounds. These include high-performance-liquid-chromatography coupled to inductively coupled plasma-mass spectrometry (HPLC-ICP-MS) and electrospray ionization-ion trap mass spectrometry (ESI-IT-MS) for the analysis of arsenic compounds, and hydride-generation gas chromatography atomic absorption spectrometry (HG-GC-AAS) for antimony compounds. Two objectives are sought in Chapter 3; one is to learn more about the pathways to the formation of arsenic compounds in mushrooms, in an attempt to address the uncertainty associated with pathways to the formation of arsenosugars and arsenobetaine, described in section 1.4.2. The other objective is to increase the knowledge of antimony behaviour in the environment by studying controlled laboratory interactions of antimony with organisms. Although some studies concerning the biological transformations of antimony have been carried out (described in section 1.5.3), clearly the number is limited, and nothing is known about the transformations of antimony in biota used for human consumption. In Chapters 4 and 5, the objective is to increase the knowledge of arsenic speciation in terrestrial ecosystems, including biota such as plants and lichens, which have not previously been studied, by using modern speciation methods. This information may be used to help determine the impact of increased arsenic loading in terrestrial environments. 20 The speciation analysis of antimony in environmental samples in Chapter 6 contributes to the understanding of antimony in the environment and adds to the limited existing data, especially for biota, as summarized in section 1.5.2. 21 References 1. Most, K . - H . Ph.D. Thesis, University of Graz, Austria, 1939; Przyoda, G . ; Feldmann, J.; Cullen, W. R. English Translation, to be published. 2. Squibb, K . S.; Fowler, B . A . In Biological and Environmental Effects of Arsenic; Fowler, B . A . , Ed . ; Elsevier: Amsterdam, 1983; pp 233-269. 3. Mervis, J. Science 1996, 273, 578. 4. McCallum, R. I. Proc. roy. Soc. Med. 1977, 70, 756-763. 5. Valentine, B . The Triumphal Chariot of Antimony; James Elliot: London, 1893. 6. Mellor, J. W. A Comprehensive Treatise on Inorganic and Theoretical Chemistry; John Wiley: N e w York, 1960; V o l . I X , pp 339-342. 7. Mortimer, C . 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Introduction Many methods have been proposed for the analysis of total arsenic in environmental samples, but only the most commonly used ones wil l be summarized here. Usually samples are digested to convert the arsenic in the sample matrix into a water soluble inorganic species. Dry ashing followed by wet digestion, and wet digestion alone are the most commonly used techniques1. Arsenic is then quantified in a sample by analyzing the solution. Photometric analysis following arsenic complexation is known 2 , but has been replaced with hydride generation (HG) techniques preceding flame atomic absorption spectrometry ( F A A S ) 3 , atomic absorption spectrometry ( A A S ) with a quartz tube atomizer 4 ' 5 ' 6, graphite furnace atomic absorption spectrometry ( G F A A S ) 7 , inductively coupled plasma (ICP) with optical emission spectrometry (OES) 8 , and ICP with mass spectrometric detection (MS) 9 . Direct analysis of solutions containing arsenic (as well as other metals) is very common, using some of the previously mentioned methods: G F A A S 1 0 , ICP-OES and I C P - M S 1 1 ' 1 2 ' 1 3 . Electrochemical methods are known as we l l 1 4 . Solid samples (subjected to little or no sample preparation, i.e., no wet digestion) are frequently analyzed by using neutron activation analysis ( N A A ) 1 5 ' 1 6 and X-ray fluorescence methods ( X R F ) 1 7 . The speciation of arsenic compounds is based almost exclusively on chromatographic methods coupled to element-specific detection. These methods include gas chromatography 28 (GC) and high performance liquid chromatography (HPLC) , as well as the less commonly used supercritical fluid chromatography (SFC) and capillary electrophoresis (CE). Arsenic compounds found in most samples are polar or ionic in character, and not volatile. Therefore they are not directly amenable to G C , so that G C methods usually involve the derivatization of the arsenic species. Hydride generation is the most common method, using sodium borohydride as the reducing agent, as shown: 3NaBHt ( a q ) + 2As(OH) 3 ( a q ) + 3 H 2 0 -> 2 A s H 3 t + 6 H 2 t + 3 N a H 2 B 0 3 ( a q ) The aqueous species As(V)/(III), C H 3 A s O ( O H ) 2 , ( C H 3 ) 2 A s O ( O H ) and ( C H 3 ) 3 A s O can be derivatized to the volatile hydrides A s H 3 , C H 3 A s H 2 , ( C H 3 ) 2 A s H and ( C H 3 ) 3 A s , respectively, with boiling points of -62.5°C, 2°C, 36°C and 52°C respectively1 8. The inorganic species of As (V) and (III) can be speciated by using different reaction conditions to form A s H 3 . More specifically, As (III) can be determined alone at pH>4 and As (III) plus As (V) can be determined quantitatively at pH<2, so that the amount of As (V) is calculated by difference. A n extraction step to selectively complex As (III) compounds can be carried out to differentiate between the normally indistinguishable (by H G ) methylarsenic (III) and methylarsenic (V) acids, which are then reacted to form their hydrides, and separated by G C 1 9 . Methylarsenic(III)thiols, but not methylarsenic (V) compounds, have been selectively derivatized into hydrides by adjusting the p H to 6 during the H G reaction and they were then detected by A A S following G C separation2 0. Most G C separations are carried out by using a packed column, but capillary G C with flame ionization detection (FED), electron capture detection (ECD) , A E S and M S , has been used as w e l l 2 1 ' 2 2 . Derivatization is carried out by using thioglycolic acid methylester 29 ( H S C H 2 C O O C H 3 ) and arsenic compounds that have been analyzed include a series of organoarsenic halides, such as Lewisite ( (ClHC=CH)AsCl 2 ) , an arsenical considered to be a chemical warfare agent. The method of H P L C can separate a range of arsenic species and does not require derivatization. The liquid sample can be directly analyzed and the results obtained may be more representative of the arsenic speciation in the original sample or extract. Arsenicals are most often identified and detected by using I C P - M S or ICP-OES for the following reasons. The flow rates from H P L C columns are similar to those used in the ICP uptake and nebulization system (around 1 ml/min) allowing for easy coupling. I C P - M S and ICP-OES are very sensitive (providing low detection limits of pg levels and ng levels, respectively) and are capable of detecting several elements simultaneously. Ion exchange chromatography is often used for the analysis of arsenic compounds, because arsenicals typically exist as anions or cations, depending on the p H used. The technique depends on the attraction between the ions and the oppositely charged stationary phase. Gel permeation chromatography has also been used for the speciation of arsenic compounds, including a trimethylarsonioriboside2 3. The use of ion-pairing chromatography is very common for arsenic speciation. In this technique, an "ion-pairing reagent", possessing an alkyl chain and an ionic end, is added to the mobile phase and allows the separation to take place on a C18 column, instead of on an ion exchange column. The current theory for ion-pairing chromatography is based on the electric double layer model. In this model, an electric double layer forms when the lipophilic end of the ion-pairing reagent interacts with the C18 column, and a counter ion interacts with the ionic end of the reagent. The resulting layers of charge enable mobility of the analyte through the column 30 to be a function of coulombic attraction, as well as interaction with the non-polar stationary phase 2 4 ' 2 5. Vesicular chromatography coupled with derivatization of the eluted analytes into their hydrides and microwave induced plasma (MIP) -AES analysis has been used for arsenic speciation 2 6. In this technique, a mobile phase containing micelles, formed from a surfactant such as didodecyldimethylammonium bromide ( D D A B ) in a concentration above the critical micellar concentration ( C M C ) , is used, together with a C I 8 column. The interaction of the analyte with the chromatographic system is thought to resemble anion exchange. Supercritical fluid chromatography (SFC) with supercritical C0 2 as the mobile phase has also been used to speciate arsenic compounds. Arsenic compounds were reacted with thioglycolic acid methylester ( T G M ) and the resulting volatile species were then extracted by using supercritical C0 2 and detected by using capillary SFC with F I D 2 7 . In another study, coupling of SFC to I C P - M S allowed separation of trimethylarsine ( (CH 3 ) 3 As) , triphenylarsine ( (C 6 H 5 ) 3 As) and triphenylarsenic oxide ( (C 6 H 5 ) 3 AsO) with no derivatization 2 8. Capillary electrophoresis (CE) has been applied to arsenic speciation. C E has been carried out with U V detection 2 9 ' 3 0 ' 3 1 , direct injection nebulizer ( D I N ) - I C P - M S 3 2 detection, and H G - I C P - M S detection 3 3 ' 3 4. The use of I C P - M S detection in the aforementioned studies gave better detection limits, but also required coupling devices that are not commercially available. The arsenic compounds As(V) , As (III), M M A A and D M A A were separated in all these studies. A study by Schwedt et al. incorporated a technique for HAs0 3S 2" speciation 3 1, which allowed the monothioarsenate ion to be determined at ppm levels in soil extracts. Mass spectrometry has been used as a detector for arsenic compounds, in some cases following H P L C separation. Conventional and microbore H P L C was coupled to liquid secondary ion mass spectrometry (LSI-MS) , continuous flow L S I - M S (CF-LSI -MS) , and direct 31 liquid introduction mass spectrometry ( D L I - M S ) to speciate arsenic animal feed additives . Fast atom bombardment ( F A B ) combined with M S as well as tandem M S ( M S / M S ) 3 6 ' 3 7 , field desorption M S ( F D - M S ) 3 6 , atmospheric pressure chemical ionization and electrospray ionization combined with a triple quadrupole M S ( A P C I - M S and E S I - M S ) 3 8 , desorption chemical ionization M S (DCI -MS) and matrix-assisted laser desorption ionization/time-of-flight M S ( M A L D I - T O F - M S ) 3 9 ' 4 0 are techniques that have been used for the characterization of arsenic compounds as standards, and in some cases, the identification of arsenic species in samples. The method of electrospray ionization (ESI) with ion trap mass spectrometry (IT-MS) was used in the present study. ESI is a soft ionization technique in which ions in solution are transformed into gas phase ions, suitable for M S analysis. It is especially useful for ionic, heat labile and high molecular weight compounds and allows M S analysis to be performed on compounds present in liquids (e.g., following H P L C separation). Ions are produced when the sample solution is nebulized and the droplets formed are subjected to a high voltage, causing their surfaces to be electrically charged. The solvent evaporates from the droplet, causing the charge density at the surface to increase; the droplet becomes unstable, breaking into smaller droplets; and sample ions are ejected into the gas phase by electrostatic repulsion from the small droplets. The ions can then be analyzed by using a mass analyzer such as an ion trap 4 1. The ion trap mass analyzer allows for high sensitivity, simplicity, and tandem techniques (MS"), among other features42. Ions are trapped in the space created by three electrodes: a ring electrode, and two end-cap electrodes. A n ac voltage of constant frequency and variable amplitude (an rf voltage) is applied to generate an electric field and its amplitude is kept at a low value initially so as to maintain stability of the ions in the trap. A mass scan takes place when the amplitude of the applied rf voltage is increased, causing instability and subsequent ejection and detection of ions of increasing m/z. M S " experiments are performed by ejecting all ions from the 32 trap except for the selected ion, and then applying a supplementary i f voltage to the end caps to translationally excite the ions. Product ions result from collisionally induced dissociation of the excited ions with helium buffer gas. 33 2.1.2. Experimental 2.1.2.1. Chemicals and reagents Arsenic standards were obtained as sodium arsenate, N a 2 H A s 0 4 . 7 H 2 0 (Aldrich), arsenic trioxide, As 2 03 (Alfa), methanearsonic acid, C H 3 A s O ( O H ) 2 (Vineland Chemical), and cacodylic acid, ( C H 3 ) 2 A s O ( O H ) (BDH) , and were dissolved in deionized water to make standard solutions. Extracts of kelp powder (Galloway's, Vancouver, B C ) and Nor i (Porphyra tenera) of known arsenosugar content 4 3 were used to identify the retention times of arsenosugars; these were verified by comparison to pure arsenosugars generously donated by K . Francesconi and T. Kaise. Arsenobetaine 4 4, arsenocholine4 5, trimethylarsine oxide 4 6 , and tetramethylarsonium iodide 4 7 had been synthesized previously according to known methods. Methanol ( H P L C grade, Fisher), tetraethylammonium hydroxide ( T E A H , 20% in water, Aldrich), malonic acid (BDH) , concentrated phosphoric acid (Aldrich), ammonium hydroxide ( I M , Fluka), pyridine (Fisher), and formic acid ( B D H ) were used as reagents for mobile phases and extractions. 2.1.2.2. Instrumentation and methods of analysis 2.1.2.2.1. HPLC-ICP-MS The H P L C apparatus consisted of a Waters 510 double piston pump, a Rheodyne six-port injection valve with a 20 pl loop, in-line filters, a guard column for each analytical column packed with the same stationary phase, and the analytical column. Columns and mobile phases are listed in Table 2.1. A V G Plasmaquad PQ2 Turbo I C P - M S ( V G Elemental) was used as a detector. Parameters for the I C P - M S are given in Table 2.2. The m/z monitored were 75, along with m/z 77 and 82 in the case of real samples, to correct for any interference from the C l - A r molecular 34 ion. j 5 C l - 4 U A r gives a m/z of 75; monitoring m/z 77 corresponding to CI- A r would allow this interference to be confirmed. However m/z 77 is also a Se isotope, and hence monitoring 8 2 Se allows differentiation from the interference. Chromatographic signals at m/z 77 were not present at the same retention time as those at m/z 75 when they appeared, and hence interference was never a problem. In most analyses, m/z 103 (Rh) was monitored as well, because R h was added to the mobile phase and used as an internal standard to correct for plasma instability. The H P L C was coupled to the spray chamber of the I C P - M S by using a minimum of P T F E tubing (10 cm x 0.5 mm id . ) with the appropriate P T F E fittings. Data from the I C P - M S were processed by using chromatographic software4 8, and identification of arsenicals in samples was made by comparison of retention times with those of standards by using at least two chromatographic systems. Semi-quantitative concentrations of arsenic compounds were determined by using external calibration curves for each compound corresponding to a matching standard, or to D M A for arsenosugars. Table 2.1. H P L C conditions for arsenic speciation Chromatography Column Mobile phase Flowrate (mL/min) Anion exchange Hamilton PRP-X100 , 150 x 4.6 or 250 x 4.6 mm 20 m M ammonium phosphate, p H 6.0 1.0 or 1.5 Cation exchange Supelcosil L C - S C X or Whatman S C X Partisil 5, 250 x 4.6 mm 20 m M pyridinium formate, p H 2.7 1.0 Ion-pairing G L Sciences ODS, 250 x 4.6 mm 10 m M T E A H , 4.5 m M malonic acid, 0.1%-0.5% M e O H , pH6 .8 0.8 35 Table 2.2. Operation parameters for I C P - M S Feature Specific Conditions Forward radio-frequency power Reflected power Cooling gas flow rate (Ar) Intermediate (auxiliary) gas flow rate (Ar) Nebulizer gas flow rate (Ar) Nebulizer type Analysis mode Quadrupole pressure <10W 0.65 L/min Time Resolved Analysis ( T R A ) 9 x 10"7 mbar de Galan 1350 W 13.8 L/min 1.002 L/min Expansion pressure 2.5 mbar 2.1.2.2.2. ESI-IT-MS Arsenosugar standards (diluted to give solutions of about 500 ppb) and the kelp extract were analyzed. The kelp was extracted according to published methods 4 3 and contained the following approximate concentrations of arsenosugars in the extract: X , 175 ppb; X I , 30 ppb; XII , 70 ppb; XIII , 220 ppb 4 9 A Finnigan L C Q ™ including a flow injection system consisting of a syringe pump delivering solutions in 50% M e O H at 100 pl/min was used. This instrument can utilize atmospheric pressure ionization (API) in two modes: ESI and atmospheric pressure chemical ionization (APCI) , o f which the former was used for this study. The ring electrode was kept at 0.76 M H z and the i f amplitude varied from 0 to 8500 V during mass scans. Experiments were carried out in negative and positive ion modes. This was done by changing the polarity of potentials applied to (a) the ion source including the electrospray capillary and the heated capillary tube; (b) the ion optics, specifically the interoctapole lens, situated between two 36 octapoles; and (c) the conversion dynode in the ion detection system. The partial pressure of helium in the ion trap was kept at 0.1 Pa (1 mTorr). For M S " experiments, the selected ions were subjected to supplementary voltages (supplied to the endcaps) ranging from 12 to 33 V and further fragmentation took place as a result of collisions with He. Trapping of fragments, fragmentation of daughter ions and changing of voltages were carried out in real time while monitoring the mass spectra. 37 2.1.3. Results and Discussion 2.1.3.1. Speciation of arsenic compounds by HPLC 2.1.3.1.1. Anion exchange chromatography The column used in these studies was a resin based column (poly(styrenedivinylbenzene)) with trimethylammonium groups providing the exchange sites (Hamilton PRP-X100 , see Table 2.1) . The elution order of some arsenic compounds when using 20 m M ammonium phosphate at p H 6 as the mobile phase is shown in Figure 2.1: arsenobetaine (AB) /As (III)/arsenosugar X , D M A , M M A , arsenosugar X I , As (V), arsenosugar X I I and arsenosugar XIII. The chromatogram obtained from a mixture of As (III), As (V), M M A and D M A was overlayed with that from a mixture of arsenosugars. Standard cationic compounds were not analyzed by using this system but they are expected to elute in the dead volume, prior to arsenobetaine. I C P - M S was used as a detector in all H P L C experiments. As (III) is present as a neutral molecule (pK a 9.3), arsenobetaine is a neutral zwitterion, and arsenosugar X is in the neutral fully protonated form. These neutral species are separable from the cationic species, and to some extent from each other. Their retention times differ by about 10 seconds; therefore, although they are not baseline resolved, i f one is present as the major species, a tentative identification can be made based on the retention time. D M A is the next eluting compound and about 50% is in the singly charged anionic form (pK a 6.28). M M A , which elutes next, is predominantly in the singly charged anionic form (98.6%, p K a i 3.6, p K a 2 8.2) . The later elution of arsenosugar X I compared with M M A may be a result of the anionic character of the phosphate group in arsenosugar X I (see Figure 1.1, structures of arsenosugars), and possibly enhanced interaction of the sugar with the resin. 38 8000 i i i i r 0 120 240 360 480 600 720 840 time(s) Figure 2.1. Chromatogram of standard arsenic compounds by using Hamilton PRP-X100 anion exchange column (15 cm) with 20 m M ammonium phosphate, p H 6. 39 A s (V) is anionic, being 90% singly charged and 10% doubly charged, hence its later elution. Arsenosugars X I I and XIII , containing sulfonate (XII) and sulfate (XIII) groups (see Figure 1.1, Chapter 1, for structures of arsenic compounds), are most likely singly charged and anionic and they elute later than As (V), when using the chromatographic system described in Figure 2.1. This later elution may be due to the increased interaction between the organic groups on these molecules with the resin. Arsenosugars have been separated on an anion exchange system in another study 5 0, where 2 0 m M ammonium carbonate at p H 10.3 was used to give the same elution order for the arsenosugars as that observed under the present conditions. The use of the carbonate mobile phase resulted in co-elution of arsenosugar X with cationic species, and co-elution of arsenosugar X I with D M A . Hence this system using ammonium phosphate as the mobile phase offers some improvement. 2.1.3.1.2. Cation exchange chromatography Cation exchange chromatography with I C P - M S detection was used to confirm or identify the compounds that co-elute in the anion exchange system described above. The chromatogram in Figure 2.2 shows the elution order of some arsenic species: arsenosugar X I , X I I I / M M A , D M A , arsenosugar X , arsenobetaine (AB) , trimethylarsine oxide ( T M A O ) , arsenocholine (AC) , and M e 4 A s + . Chromatographic peaks corresponding to the arsenic species A s (V) and As (III) are not shown in Figure 2.2, to provide clarity of presentation, since these compounds elute closely together with other species. Nevertheless, their retention behaviour was observed and is discussed below. The chromatograms for a mixture of standards ( M M A , A B , T M A O , A C , and M e 4 A s + ) was overlayed with the chromatogram for kelp extract, containing arsenosugars X , X I , X I I (with unknown retention time) and XIII . 40 35000 </> 30000 H 25000 -20000 15000 10000 5000 0 standard cationic species kelp extract I I I I l i 0 180 360 540 720 900 1080 time (s) Figure 2.2. Chromatogram of standard arsenic compounds by using Whatman SCX cation exchange column with 20 mM pyridinium formate, pH 2.7. 41 A silica gel based column containing benzene sulfonic acid functional groups as exchange sites (Whatman S C X or Supelcosil L C - S C X , see Table 2.1) was used with a mobile phase consisting of 20 m M pyridinium formate at p H 2.7, based on methods described in previous studies 5 1 ' 5 2. A t this pH, As (V) has 74% singly charged anionic character and is unretained by the column (not shown in Figure 4). p K a ' s for the individual arsenosugars are not known but the p K a for the dimethylarsinoyl moiety on the sugars is estimated to be 3.85 5 3 , indicating that the arsenosugars should be somewhat cationic at this pH. However, the co-elution of arsenosugar X I with As (V) (not shown in Figure 2.2), which is unretained, indicates that arsenosugar X I may be more anionic or neutral in character. Arsenosugar XIII, on the other hand, was observed to co-elute with the neutral species As (III) and M M A (As (III) not shown in Figure 2.2). D M A is separated from the neutral species, indicating some cationic behaviour, possibly as M e 2 A s + ( O H ) 2 5 4 . The partial protonation of the oxygen in the dimethylarsinoyl moiety in arsenosugar X would give it some cationic character, allowing it to be retained on the column. Arsenobetaine has a p K a o f 2.18 5 4 and is 20% cationic in nature (80% zwitterionic and neutral at this pH). It is probably more cationic than the previous two species, and hence retained longer on the column. T M A O is thought to exist as [ ( M e 3 A s O H ) + (OH)"], from the hydrolysis of M e 3 A s 0 5 5 , although the peak is very broad under these conditions, which may indicate the presence of more than one species. Arsenocholine and M e 4 A s + are cations irrespective of pH, and their retention behaviour under these conditions indicate that arsenocholine is less strongly retained. Previous studies have shown the same elution order for these two compounds under the same conditions, but the elution order was reversed when a bare silica column was used 5 4, and when an ion-pairing system with a reversed phase column and sulfonate mobile phase was used 5 5. N o explanation was given in the former study, and in the latter study arsenocholine was suggested to be more hydrophobic. 42 The organic nature of the pyridinium mobile phase may cause the more hydrophobic arsenocholine to be eluted faster from the cation exchange column in the present study. However, other interactions, such as hydrogen bonding between silica O H groups and the O H group on arsenocholine, must be responsible for its longer retention time on the bare silica column. 2.1.3.1.3. Ion-pairing chromatography When the presence of arsenosugars is indicated by one or both of the two previously mentioned chromatographic systems, it can be confirmed by using a third chromatographic system. The system used here is one that has been developed for the analysis of anions, particularly arsenosugars56. Ion-pairing chromatography with I C P - M S detection is used, a technique that combines a mobile phase containing an ion-pairing reagent with a reversed phase column. Tetraethylammonium hydroxide ( T E A H ) is the ion-pairing reagent, and the mobile phase is adjusted to a p H of 6.8 with malonic acid and nitric acid. A small amount of methanol is added (0.1-0.5% v/v). Methanol can be used to control chromatographic behaviour in reversed phase systems, with increased concentrations resulting in shorter retention times for the arsenosugars. Methanol has also been found to increase sensitivity of arsenic compounds when using I C P - M S detection 5 7. The elution order shown in Figure 2.3 for some arsenic species is arsenobetaine (AB) , M M A , D M A / A s (V), arsenosugar X , X I / X I I , and XIII . Chromatograms of A B , M M A , D M A and As (V) standards were overlayed with kelp extract and also with Nor i extract. Nor i extract is known to contain arsenosugars X and X I as the only arsenosugars43. 25000 Nori extract standards and kelp extract 20000 - x , XIII A B M M A I I I X 15000 - I w i 0 240 480 720 960 1200 time (s) Figure 2.3. Chromatogram of standard arsenic compounds by using ion-pairing reversed phase chromatography, with G L Sciences C I 8 column and 10 m M TEAH/4 .5 m M malonic acid, p H 6.8, 0.1% M e O H . 44 A s (III) elutes closely after arsenobetaine and is riot'shown on Figure 2.3 to retain clarity. Arsenobetaine and A s (III) are expected to be neutral molecules at p H 6.8, and to have short retention times. M M A and D M A elute in the opposite order compared to anion exchange; this may be due to enhanced interaction of D M A (containing 2 methyl groups) with the C18 chains of the stationary phase. D M A co-elutes with A s (V), which was also seen by other researchers58. Using the present chromatographic system, the arsenosugars are separated from the other ions, although arsenosugars X I and X I I are not baseline resolved from each other. Figure 2.3 shows that arsenosugar X I alone (dotted line) elutes slightly later, which allows this compound to be identified in the absence of arsenosugar XII . Likewise, arsenosugar X I I can be identified in the absence of arsenosugar X I . The elution order for the sugars most likely indicates increasing anionic character when going from arsenosugar X to XIII. Although not one of these chromatographic systems can be used alone for the analysis of complex mixtures, such as those found in environmental samples, a combination of anion, cation and ion-pairing chromatography can be used to obtain a satisfactory separation and identification of at least 11 arsenic species. Analyzing a sample by using two or three different chromatographic systems, although time consuming for routine use, is a very useful tool for strengthening the identification of arsenic compounds. 2.1.3.2. Characterization of arsenosugars and compounds in kelp powder extract Kelp powder has been previously analyzed and it has been suggested that this sample contains the four arsenosugars X , X I , XI I , and XIII, based on H P L C retention times 4 3 ' 4 9 . As a result of this identification, kelp extract is often used as a reference for the retention times of these arsenosugars, and the retention times are then used to identify arsenosugars in other 45 samples of unknown arsenic speciation. A n attempt was made to identify the arsenosugars in kelp extract by using a mild ionization mass spectrometric technique, possible with the Finnigan L C Q ™ E S I - I T - M S instrument, as well as by analyzing pure arsenosugar standards. Such identification would validate other identifications based on retention times of the reference sample. The positive-ion ionspray (IS) tandem mass spectra (i.e., M S - M S ) of pure arsenosugars X , X I , X I I and XII I have been presented by others 5 9. Positive and negative-ion fast atom bombardment ( F A B ) tandem mass spectra have been produced for arsenosugars X I , X I I and XIII , as we l l 6 0 . It was of interest to compare the fragmentation behaviour of arsenosugars in the present M S - M S experiments by using low-energy, low pressure ( lmTorr) C I D conditions in the ion trap, with that observed by using low-energy, higher pressure (4-18mTorr) 6 1 C ID M S - M S conditions (IS with a triple-quadrupole mass filter). Because Corr et al.59 observed only positive ions, comparisons were also made between the present results and the results obtained by Pergantis etal.60 (high-energy C I D conditions, with F A B reverse-geometry four-sector mass analyzer) for negative and positive-ion mass spectra. A summary of M S and M S - M S analyses obtained from pure arsenosugars is given in Table 2.3. 46 Table 2.3. ES I - IT -MS experiments and fragments for pure arsenosugars. Arsenosugar Precursor ion m/z Product ion m/z Relative abundance (%) (mode) selected X, MW = 328 329 329 46 (positive) 311 52 ' 237 100 195 26 329 -> 311 311 100 209 30 329 -> 237 237 100 219 75 XI, MW = 482 483 391 65 (positive) 329 100 237 13 483 -» 329 329 100 237 55 XI, MW = 482 481 389 100 (negative) 245 27 481 ->389 389 100 267 20 223 49 193 33 XII, MW = 392 393 392 100 (positive) 281 53 XII, MW = 392 391 391 52 (negative) 373 100 269 48 391 ->269 269 100 225 90 197 17 XIII, MW = 408 407 408 ' 24 (negative) 389 100 285 62 171 52 407 -> 389 171 100 407 -> 285 285 21 241 100 213 35 153 75 137 28 97 27 87 30 407 ->• 389 -> 171 171 35 97 100 47 Only low concentrations of arsenosugars were available and the mass spectrometer was contaminated by previous samples, which caused high background levels during the arsenosugar analyses. Hence, the M S " capability of the ion trap mass analyzer in the mass spectrometer was useful because it allowed isolation of specific ions of the sugars and subsequent fragmentation. Only the two sugars X I and X I I could be analyzed by both negative and positive modes to give useful information. Molecular type ions were not observed in large enough abundance for trapping and subsequent M S - M S analysis for the other two sugars, X and XIII , in the negative and positive modes, respectively. A s mentioned earlier, arsenosugar X ( M W = 328) was characterized in the positive ionization mode by Corr et al.59 by using an ionspray source and triple quadrupole mass spectrometer with M S - M S capabilities. They found a single product ion forming at m/z 237, corresponding to the fragment containing arsenic and the ribose group. Other previous studies showed the formation of fragments at m/z 311, 221 and 177 for this compound 4 7. In the present work (Figure 2.4), the fragments at m/z 311 (loss of H 2 0 ) , 237 (with the structure shown in Figure 2.4) and 209 are produced when the (M+H) + ion at m/z 329 is trapped and subsequently fragmented. The fragments obtained when m/z 311 ([ (M+H)-H 2 0] + ) was trapped and fragmented are m/z 209 and 195, which have not yet been identified. The product ion obtained when m/z 237 was trapped and fragmented is m/z 219 (resulting from loss of water). Hence these results show similarity to both of the previous studies, where the fragment of m/z 237 is common with the study by Corr et al.59 and the fragment of m/z 311 is common with the study by Cullen et al.47 Although Pergantis et al.60 did not analyze standard arsenosugar X , they were able to isolate a precursor ion at m/z 329 in the positive-ion mode from an algal extract, and found fragments at m/z 311 and 237 (similar to the present study), as well as at m/z 208, 176, 165, 122 and 97. 48 Both positive and negative modes of ionization were successful for arsenosugar X I ( M W = 482), as mentioned before. When (M+H) + (m/z 483) was trapped and fragmented (see Figure 2.5), m/z 391 corresponding to loss of ( O H ) C H 2 ( C H O H ) ( C H 2 O H ) is observed. A fragment of m/z 329, corresponding to arsenosugar X is observed, as well as m/z 237, corresponding to the dimethylarsenic-ribose moiety. These fragments were also observed in the previous studies by Corr et al.59 and Pergantis et al60. The higher energy C I D conditions led to additional ions in those studies. When arsenosugar X I was analyzed in the negative ionization mode, and (M-H)" was trapped and fragmented (see Table 2.3 for these results, and Figure 2.6 for the structure of ( M -H)"), m/z 389 is observed as the negative ion analogous to the ion of m/z 391 described above. A product ion of m/z 245 is also observed, corresponding to the fragment remaining when the dimethylarsenic-ribose moiety is lost, with the structure [ ( C H 2 C H ( O H ) C H 2 O H ) 2 P C V ] . When m/z 389 was trapped and fragmented (Figure 2.6), m/z 267 is produced, having the structure shown in Figure 2.6, after loss of the dimethylarsenic group. The fragment at m/z 223 may indicate loss of C H 2 = C H O H (44 amu) from the m/z 267 fragment. The fragment at m/z 193 is unidentified. Pergantis et al.60 observed m/z 389 and 245 as well, but not 267 or 223. 50 O) -0 0 CO O) co ro oo r • co cn O -r-co : o t CO O) $ ~ O I ro iri —L co t ug I"5. . o -co C M p CM | to CM ; o I CM oi ; CM -in CM if) CM O CM O O) CM CM h8 CM CM O) — f P, t CM o> =i-o ro -CM: © o IT) O IT) CD o © in IT) • f l -ic) CM o CM o eouepunqv aA»e|ay u • O o e (D I OA c O N oo m N C <D o (/3 O O N % % s I c/3 I-H X oo O C (D B o S o u a. bo s 2 & 5 2 Arsenosugar X I I ( M W = 392) was successfully fragmented in both the positive and negative ionization modes. In the positive mode, trapping of m/z 393 (M+H + ) ion and M S - M S analysis (Figure 2.7) gives a large abundance of m/z 392 (M» + ) and no m/z 393 is observed. A fragment having m/z 281, which may correspond to loss of C H 2 - S 0 3 H and O H (see Figure 2.7) is also observed. These ions (m/z 393 and 281) were not observed when this compound was subjected to ionspray M S in the positive mode by Corr et al.59 or to F A B - M S by Pergantis et al.60 Those authors observed m/z 375, 296 and 237 5 9 , and 375, 237, 165 and 122 6 0 , fragments that were found only in very small abundance or not at all in the present study. More structural information is obtained when arsenosugar X I I was analyzed in the negative mode. The [M-H]" ion of m/z 391 was trapped and fragmented (Figure 2.8) and loss of H 2 0 gives m/z 373, and the dimethylarsenic moiety (m/z 269, see Figure 2.8 for structure) is lost. When m/z 269 was subsequently trapped and fragmented, loss of C H 2 = C H O H (269-44) followed by loss of C O (225-28) may have occurred to form the fragments at m/z 225 and 197 (structures ( C = 0 ) C H ( O H ) C H O R and (OH)CH=CH(OR), where R is C H 2 C H ( O H ) C H 2 S 0 3 " ) . Interestingly, m/z 197, but not m/z 225 was observed by Pergantis et al.60; m/z 197 was assigned a similar structure to the one proposed in the present study. Arsenosugar XIII ( M W = 408) was ionized successfully only in the negative mode, indicating greater stability of the molecule in the anionic sulfate form compared to its stability in the cationic protonated form. This enhanced stability of the anion corresponds well to the more anionic character of this sugar as observed in its H P L C behaviour in the previous section. The anion (M-H)" at m/z 407 was trapped and fragmented (Figure 2.9) to produce the following ions: m/z 389, corresponding to loss of H 2 0 ; m/z 285, corresponding to loss of the dimethylarsenic moiety as described above; and m/z 171, corresponding to [ H O - C H 2 - ( C H O H ) - C H 2 - 0 - S 0 3 " ] (i.e., loss of the dimethylarsenic-ribose moiety). Subsequent trapping of m/z 389 produced the 53 aforementioned fragment at m/z 171. Trapping of m/z 285, (Figure 2.10) produced fragments at m/z 241 and 213, which seem to follow the same pattern as the fragments at m/z 225 and 197 for the ion at m/z 269 in arsenosugar XI I , which may again indicate loss of CH2=CHOH and then C O . Other product ions from m/z 285 are found at m/z 153 (CH 2 =C(OH)-CH 2 -0 -S0 3 " ) , 137 (not easy to identify, but possibly resulting from loss of O from m/z 153), 97 (HO-SO3") and 87 (unidentified). The fragmentation of m/z 171 produced m/z 97 (HO-SO3"). Pergantis et al60 observed some of the same fragments: m/z 389, 285, 213 (but not 241), 171, 153 and 97. Useful structural information was obtained in the negative ion mode for all sugars but arsenosugar X . The loss of the 122 amu fragment was observed for sugars X I , X I I and XIII in the negative mode and can be considered diagnostic of this group of compounds 6 0. In general, fewer fragments were observed in the positive mode of analysis. Characteristic arsenic containing fragments were observed in arsenosugars X , X I and X I I in the positive mode, however. The previous studies reported a greater number of fragments compared with those in the present study, resulting from the use of higher energy CUD conditions in those studies. M S -M S - M S experiments were conducted in the present study, providing new information about the behaviour of some of the arsenosugars in this mode. 54 o C O -f-co CM : co j O) S o co; co r- i C M t co i-f o r-to C M : co m : co • •a o e > s o *«3 O N cn 5 T £ o 5-15 X 8 .AT1 \ X o (0 CM C O r O • C O (-CM 1- ; O O r-o co • co T ; O o r-«D in i CM C M » " r. C O -m : C M r •s '% s I s 1 TJ C ^> p X t * c 3 60 3 no O C <D Vi t-03 C M If i -C M C M C O t C M ; O o o m co o oo in CO m m o in in eouepunqv aAgeisy T t o o t/3 00 fa 56 100-389.0 95-9 0 : 85^ 80^ 7 5 : 70 : 65-60-g 55-to T 3 C | 50-> 1 45^ a: 40-35-30-25H 20-15-10-5-0-O H O H 285.3 171.4 153.3 137.3 129.4 | 1 213.3 2 4 ^ - 3 275.3 199.3 200 229.3 271.3 >50~ 286.6 314.3 345.2 363.3 300 ' m/z l — r i i 350 408.0 400 408.9 Figure 2.9. Mass spectrum of arsenosugar XIII standard, MS-MS with m/z 407 selected, negative mode. 5 7 100-95: 90-85H 241. 80-75H 70-65 60-c 55^  ro £Z 3 J < 50H 1 45J tr: j 40-35-30-2 0 : 10: 5-0 87.1 153.1 97.1 137.1 213.1 171.1 123.1 100 195.1 150 200 285.2 2407 241.9 250"~ 302.5 - i — i — r 359.4 j5o J50 75o~ m/z Figure 2.10. Mass spectrum of arsenosugar XIII standard, M S - M S - M S with m/z 407 selected, then m/z 285, negative mode. 58 A fUll mass scan in both the positive and negative modes of the crude kelp powder extract did not show the arsenosugars as major ions. However, the M S " capability of the L C Q ™ instrument proved valuable since the ions characteristic of the individual arsenosugars (m/z 329 (positive mode) for arsenosugar X , m/z 481 (negative mode) for arsenosugar X I , m/z 391 (negative mode) for arsenosugar X I I and m/z 407 (negative mode) for arsenosugar XIII) could be selected from the complex matrix and then fragmented to ascertain i f the characteristic product ions were observed. Some of the fragments that were produced during the M S and M S - M S experiments are summarized in Table 2.4. The fragments listed are ones that also appear in the pure arsenosugar standards. 59 Table 2.4. ESI - IT -MS experiments for kelp powder extract Arsenosugar Precursor ion m/z Product ion m/z Relative abundance (mode) selected (%) X , M W = 328 329 329 100 (positive) 328 82 311 7 237 11 X I , M W = 482 481 481 17 (negative) 463 100 389 30 245 27 XII , M W = 392 391 391 100 (negative) 373 85 225 6 XIII, M W = 408 407 407 80 (negative) 389 100 285 30 407 -> 285 285 100 241 25 The fragments that might be considered characteristic occur in very low abundance and in some cases are no more abundant than the other peaks that appear in the mass spectra. This is the case for the precursor ion of m/z 329 (arsenosugar X ) and m/z 391 (arsenosugar XII); the characteristic m/z 237 for arsenosugar X , and m/z 225 for arsenosugar X I I have abundances of 11 and 6%, respectively (Table 2.4). Another problem is that other fragments are observed in addition to those obtained from M S - M S experiments with standard compounds and these are probably due to the impurity of the 60 sample (i.e., precursor ions not with the structure of the arsenosugar may be trapped). Interestingly, some of the other fragments that occur when m/z 481 is trapped and dissociated in the negative mode (Figure 2.11) were also observed by Pergantis et al.60 (e.g., m/z 407 and 287), but not when the standard compound was analyzed in the present study. The ion occurring at 100% abundance, m/z 463, indicates loss of water, but it is also not observed for the standard. Again, the characteristic ions at m/z 389 and 245 are at an abundance not significantly different from the other fragments whose structures are unknown. L o w concentrations for arsenosugars X I and X I I (about 15 and 35 ppb, respectively, after 1:1 dilution of the extract with methanol), made further confirmation by M S - M S - M S experiments impossible. In the positive mode, only the (M+H) + ion (m/z 329) for arsenosugar X could be isolated from the complex mixture, and in addition to the low abundance of the characteristic m/z 237, the presence of M « + ion at m/z 328, a fragment that was not observed previously, can be seen (Table 2.4). When the (M-H)" ion for arsenosugar XIII in the kelp extract (m/z 407) was trapped, the characteristic fragments of m/z 389 (loss of water) and 285 (loss of dimethylarsenic group) are observed. When m/z 285 was subsequently trapped, the fragment of m/z 241, observed in the same experiment with the standard, is seen (Figure 2.12). Another fragment of appreciable abundance is present in Figure 2.12a at m/z 363, indicating either loss of 44 amu ( C H 2 = C H O H or C 0 2 ) from m/z 407, or loss of 26 amu ( C 2 H 2 ) from m/z 389. This fragment was not observed in the mass spectra generated for the standard compound, nor in any other reports. The mechanisms are not obvious for the suggested mass losses and they may involve ring cleavage or molecular rearrangement. However, a similar mass loss is observed from m/z 481/463 (sugar XI) and 391/373 (sugar XII) . Dissociation processes are apparently taking place that are specific to these compounds in the kelp extract. 61 6 2 o c o o c 53 i ! o M O ! co T-c\i —1 CO CM -A • Irt asuepunqv aNjeiaa CM O ) -oo co oo oo ro co-co ro oo I o_L CM i co i CM -CD i n rg o r*o vo ! io oo l C M • L O - L cn oo —r I o CO CM CM to o> L O oo o LO CO o CO o L O L O o aouepunqv aAijeisy & . f a o a a •§ t> 5 Q ^ u a O .53 -e 3 CN u c cd eu L _ , X t » 3 1 N ja •s a C ^ « « N ^ 11 L . CD 3 60 a 3 o *n <+* oo ccj <N is N c/i C fa ^ fa B 63 The strongest argument for the presence of arsenosugars in this extract is the evidence for arsenosugar XIII because the M S - M S , as well as M S - M S - M S experiments, show characteristic fragments. The identification of the other compounds, based on this data alone, is not as clear. The major reason for the problems with compound identification in the kelp extract is the low levels of the compounds, especially in the presence of large amounts of matrix components. Chromatographic separation followed by these M S experiments would improve the identification of the arsenosugars. These results are very useful in characterizing the arsenosugar standards, verifying the cationic behaviour of arsenosugar X , the similar anionic behaviours of arsenosugars X I and XII , and the strongly anionic character of arsenosugar XIII , which was also observed in the H P L C behaviour of these compounds. Moreover, this mass spectral information adds to that currently available in the literature. 64 2.2. Methods for the Analysis of Antimony 2.2.1. Introduction Methods for the analysis of total antimony in samples are similar to those used for arsenic, which are described briefly in section 2.1.1. Speciation techniques for antimony are fewer in number. A recent review lists the known methods for speciation, including extraction techniques; electrochemical techniques; coupled techniques such as hydride generation-gas chromatography ( H G - G C ) with flame-in-tube A A S or I C P - M S detection; and high-performance liquid chromatography ( H P L C ) with ICP-OES, ICP-M S or H G - A A S detection 6 2. O f these, only H G - G C - A A S , H G - G C - M S and H P L C - I C P - M S methods were used for this work. H P L C techniques are well established for the speciation of arsenic compounds (section 2.1.1), but only recently have H P L C methods been developed to speciate antimony. In the first report of such methods, ICP-OES detection and a cation exchange column were used to separate Sb (III) and Sb (V) in electrolyte solutions 6 3. Anion exchange techniques coupled to I C P - M S or ICP-OES were explored by other authors, where mobile phases such as tartrate at p H 5.5 6 4 and phthalate at p H 5 6 5 ' 6 6 ' 6 7 , 2 m M K O H and E D T A at p H 4.5 6 8 were used successfully to separate Sb (III) and Sb (V) , and M e 3 S b C l 2 or M e 3 S b 0 6 7 ' 6 8 . These methods have been used with some success to analyze real environmental samples 6 5 ' 6 7 ' 6 8. Some of these methods were also attempted for the separation of dimethylantimony compounds of unknown structure but with no 69 success . The application of H P L C to antimony speciation is limited by the lack of standard compounds available; for example, no standard dimethyl and monomethylantimony (V) compounds are commercially available, or easily synthesized. A t the pHs studied for most of the 65 H P L C method development, Sb (V) exists as the anion Sb(OH)6_ whereas Sb (III) exists as Sb(OH) 3 when the oxide is dissolved, or as a complexed form such as the antimonyl tartrate, [Sb 2 (C 4 06H 2 ) 2 ] 2 ~. The Sb (V) anion is not strongly retained on an anion exchange column and is hence easily eluted from an anion exchange H P L C column. Uncomplexed Sb (III), however, precipitates easily as the oxide, and in the complexed anionic form is strongly retained on an anion exchange column. M e 3 S b C l 2 most likely exists at neutral p H as M e 3 S b ( O H ) 2 and is not retained or is retained to a minor extent on an anion exchange column 6 7 ' 6 8 . As described in section 2.1.1 for arsenic, H G - G C speciation techniques can be used to detect antimony species in samples as inorganic antimony compounds in the +3 and +5 oxidation states, which are derivatized to form SbH 3 ' ' . It is also be used for compounds that can be derivatized to form methyl-, dimethyl- and trimethylstibines (MeSbH 2 , M e 2 S b H and M e 3 S b ) 7 0 ' 7 3 . A problem has been noted by various researchers attempting to apply the hydride generation reaction to the generation of methylated stibines: when trimethylantimony di chloride (Me 3 SbCl 2 ) , trimethylantimony dihydroxide (Me 3 Sb(OH) 2 ) and dimethylantimony dihyroperoxychloride (Me 2 SbCl (0 2 H) 2 ) were reacted with borohydride and acid to form their corresponding stibine, four peaks corresponding to SbH 3 , M e S b H 2 , M e 2 S b H , and Me 3 Sb, appeared for each compound, rather than the anticipated ones of M e 3 S b or M e 2 S b H 7 3 ' 7 4 Dodd et al13 postulated that this "rearrangement" occurred when the reaction apparatus had not been conditioned properly and found that the problem disappeared when the apparatus was rinsed with the reagents ( N a B I L solution and acid solution) for three minutes or more. In the following section, some problems with the hydride generation derivatization technique will be described for antimony, including examples of rearrangement/ demethylation which can lead to misinterpretation of analytical results. 66 2.2.2. Experimental 2.2.2A. Chemicals and reagents Antimony (V) and (III) standards were obtained as potassium hexahydroxyantimonate, K S b ( O H ) 6 (Aldrich), and potassium antimonyl tartrate, K2Sb2(C406H2)2 (Aldrich). M e 3 S b C l 2 was synthesized as described elsewhere7 5. Stock solutions were made by dissolving these compounds in deionized water and diluting the resulting solutions to 1000 or 100 mg L" 1 as Sb. Standard working solutions were made by diluting the stock solution with deionized water as necessary. NaBFLi (reagent grade, Aldrich) was dissolved in deionized water fresh daily to provide a concentration of 2% w/v. Glacial acetic acid, citric acid, sodium hydroxide (for p H adjustment), maleic acid and concentrated sulfuric acid were all reagent grade and obtained from common distributors. 2.2.2.2. Method of analysis for HG-GC-AAS Three methods were used: Methods 1, 2 and 3. The apparatus for Methods 1 and 2 was composed of a semi-continuous flow, hydride generation system developed for arsenic analysis7 6, coupled to an atomic absorption spectrometer (Varian AA1275) fitted with an Sb lamp (Varian) operating at a wavelength of 217.6 nm. One modification was made to the basic apparatus in the form of using a gas-liquid separator7 7 that resulted in less analyte carryover. The apparatus consisted of Tygon tubing for the peristaltic pump, and P T F E tubing (1/8" OD) for the remainder. The glass gas-liquid separator was silanized with ( C H 3 ) 2 S i C l 2 before use. For Method 3 the semi-continuous flow and batch modes of analysis were combined. Figure 2.13 shows a schematic diagram of the apparatus used. Unsilanized glass batch reactors 67 of 60 ml volume were incorporated into the apparatus from Method 1. A gas-liquid separator was not used for Method 3 since the gases were separated from the liquids in the batch reactors. The A A S , peristaltic pump and tubing were identical to those used in Method 1. For all methods, data were collected from the A A S and processed directly by using an H P 3 3 90A integrator, or were analyzed with the aid of Shimadzu EZChrom software. For Method 1, a peristaltic pump was used to deliver standard or sample solution (ranging from 5 pJL to 200 u,L for standards, and from 1 mL to 5 mL for samples) to mix with the acid or buffer and then to mix with a solution of NaBEL, (2% w/v) in a reaction coil. For Method 2, standard solution was mixed with a solution of NaBIL, (2% w/v) in the mixing coil, and the gas-liquid mixture was mixed with I M H 2 S 0 4 . For both Methods 1 and 2, the gases evolved were separated in the gas-liquid separator and then swept by a flow of helium through a P T F E U-tube at -78 °C (dry ice/acetone) to remove water and into a P T F E U-tube, where they were trapped at -196 °C (liquid N 2 ) . Continuous hydride generation and trapping were carried out for 3 minutes. The peristaltic pump was then stopped (making the system semi-continuous) and the second U-tube was heated to 60 °C, allowing the gases to be swept with He at a flow rate of 40 mL/min onto a Poropak PS column, which was then heated from 70 °C to 150 °C at a rate of 30 °C/min, whereby the gases were separated. They were then detected by A A S . 68 69 For Method 3, the peristaltic pump delivered standard solution and 2% (w/v) NaBFL, solution to the first batch reaction vessel (A). A volume of 30 mL was delivered to reactor A while helium gas was bubbled through the glass frit to strip the solution of gases. The gas stream was bubbled through the contents of reactor B (see Figure 2.13). Throughout the process, the gas stream was dried and trapped at -196 °C and the remaining procedure was identical to the one previously described for Methods 1 and 2. Reactants were added to reactor B in Method 3 from a 1 m L syringe inserted between the rubber stopper and the side of the reactor. Measurement of p H was carried out with an Accumet Model 15 p H meter (Fisher Scientific) after the sample and acid had been mixed, but before the NaBFL, was added. 2.2.2.3. Sample preparation Mycelia of the pink oyster mushroom Pleurotus flabellatus (Western Biologicals, Aldergrove, B C ) were grown with shaking in 400 mL potato dextrose broth (Difco) in a 1 L Erlenmeyer flask. Me3SbCl2 solution was added to the broth to give a concentration of 1 ppm in antimony. After a 14 day growing period, the mycelia, as spherical pellets, were harvested by centrifugation and rinsed with distilled water. They were then homogenized with an Ultraturrax T25 homogenizer (Jak & Kunkel) to give a solution of lysed fungal cells, and analyzed by using H G - G C - A A S . A control experiment was carried out in which the fungus was grown in the same manner, only without the addition of antimony to the potato dextrose broth. See Chapter 3 for more details about this experiment. 70 2.2.3. Results and Discussion 2.2.3.1. Demethylation of trimethylantimony species in aqueous solution during analysis by using G C - H G - A A S Dodd et al73, observed that peaks corresponding to SbH-3, M e S b H 2 , and Me2SbH, as well as the expected Me 3 Sb , appeared when the hydride generation apparatus had not been preconditioned with the reagents used for analysis (2% N a B H j and 4 M acetic acid). In an attempt to replicate this observation, experiments were carried out by using the same method, but with an A A S instead of an M S as the detector. M e 3 S b C l 2 was analyzed after a preconditioning step which consisted of rinsing all tubing with water, as well as after rinsing with 2% NaBEL, and 4 M acetic acid. In all replicates (five for each), the results were the same when Me3SbCl2 was reduced and analyzed: M e 3 S b and a minor amount of M e 2 S b H (corresponding to <2% demethylation) were detected. The large amount of demethylation that was observed by Dodd et al.73 could not be replicated. 2.2.3.1.1. The effect of acid When HC1 was used as the acid in the reaction to adjust the p H of the sample, increased demethylation was observed. Different concentrations of HC1 led to the appearance of SbH 3 , M e S b H 2 and M e 2 S b H in differing amounts, indicating a p H dependence for the demethylation process. Hence, demethylation of M e 3 S b C l 2 at different acidities was studied, and the results obtained are illustrated in Figure 2.14. It was assumed that the reaction efficiency and detector response for each species is the same (which was observed to be the case for M e 3 S b and SbH 3 ) , allowing the amount of each stibine to be determined. A minimum of three replicates were carried out at each pH, but reproducibility of the normalized amounts was poor, with relative 71 100 90 80 70 15 £ 10 H 5 H T - CO m r--CO CO B co CO I T o a> • * T -CN PH IT) CM oi O O CM CO t - CN CO CO CO co (6 Figure 2.14. Percent amounts of stibines generated from M e 3 S b C l 2 at varying pH, when using H G - G C - A A S . Amounts of stibines may not add up to 100% because of imprecision in blank correction. Except where indicated, unbuffered HC1 was used to adjust the pH. A = sulfuric acid, B = maleic acid, C = citric acid, D = citrate buffer, E = water. 72 standard deviations averaging 22% (0.1% up to 50% for amounts approaching the detection limit). The acid used in most of the experiments was unbuffered HC1, but at some pHs different buffers and acids were tested to determine i f the demethylation was specific to HC1. These buffers and acids are specified in Figure 2.14. Small amounts of M e 2 S b H appear at p H 3.22 and p H 3.30, but at p H 6.10 no demethylation is seen for the concentration of M e 3 S b C l 2 being studied. Although a statistically significant dependence of demethylation on the acid used was not observed for the acid concentrations studied (see Figure 2.14), it must be noted that certain acid systems give less demethylation than would be expected considering the pH. For example, reactions carried out by using 4 M acetic acid (pH 2.2) resulted in statistically significantly less demethylation than experiments carried out at similar pH, by using other acids (see Table 2.5 and compare to results for 0.01 M HC1 and 0.1 M citric acid). However, when 0.6 M acetic acid was tested (Table 2.5), the amount of demethylation was not statistically significantly different from other acid systems at similar pH, but the amount of demethylation was calculated to be statistically significantly different from that observed when 4 M acetic acid was used. More concentrated solutions of citric acid (0.5 M and 1 M ) also showed amounts of demethylation that were calculated to be statically significantly less than that observed when 0.1 M citric acid was used (Table 2.5). More studies must be carried out to elucidate the mechanism of the demethylation phenomenon and hence to explain this observed behaviour. 73 Table 2.5. Comparison of amounts of demethylation when using different concentrations of acids. Lower values of percent Me 3 Sb of total Sb indicate higher amounts of demethylation. Acid used p H (calculated)3 % M e 3 S b of total Sb (SD C) 0.01 M H C 1 2.25 b 92.5 (0.6) 0.6 M acetic acid 2.6 94 (2) 4 M acetic acid 2.2 98 (2) 0.1 M citric acid 2.19 b, 2.2 92.2 (0.7) 0.5 M citric acid 1.9 96.6 (0.6) 1 M citric acid 1.7 95.9 (0.9) a Unless otherwise stated, pH is calculated by using pH=-log(VKa x [HA]/2); where K a = 1.75 x 10"5 for acetic acid; K a = 7.44 x 10"4 for citric acid; [HA] = concentration of acid. [HA] is divided by 2 because of dilution during mixing of the acid in the HG apparatus. b pH was measured after mixing the acid solution with water, as described in section 2.2.2.2. c SD = standard deviation, calculated from 3 replicate analyses. 2.2.3.1.2. The effect of concentration When high levels of M e 3 S b C l 2 (500 - 1000 ng) were analyzed, demethylation was observed at neutral p H as well. Demethylation can only be seen when high levels of M e 3 S b C l 2 are analyzed, even though the amount of demethylation is low at neutral p H (<1%), because the levels of SbH 3 , M e S b H 2 and M e 2 S b H are sufficiently high for detection. The detection limits are estimated to be between 1 ng and 5 ng for all the stibines, although standards are not available for methyl- and dimethylantimony species. The demethylation pattern is not dependent on the concentration of antimony being analyzed. Small ranges of concentration (2 orders of magnitude) were tested, however, so the possibility that the demethylation pattern changes at higher concentrations cannot be discounted. 74 2.2.3.1.3. The effect of sample matrix To illustrate the importance of establishing the presence of demethylation during the analysis of trimethylantimony species, a sample known to contain trimethylantimony species was analyzed. We found that the sample matrix in this example has an effect on the hydride generation of methylstibines. The term "sample matrix" refers to any chemical components other than the analytes (antimony species) that make up the sample. Figure 2.15a shows a chromatogram of an aqueous extract of the fungus Pleurotus flabellatus that had been grown in liquid culture and amended with M e 3 S b C l 2 . The stibines were generated with the aid of aqueous borohydride with no acid or buffer, conditions under which the M e 3 S b C l 2 standard in water described in section 2.2.3.1.1 showed minimum demethylation (pH 5.6). Figure 2.15a shows peaks in addition to the one corresponding to the expected trimethylstibine for this sample. Figure 2.15b shows the same fungus extract analyzed while using a 0 .05M citrate buffer at p H 6.2. Most of the demethylation appearing in Figure 2.15a was eliminated by using a buffer. To determine whether the demethylation observed in Figure 2.15a was caused by the sample matrix, a control sample from the mushroom culture that had not been amended with antimony, and hence with the identical matrix but found previously to contain a concentration of antimony less than 0.05 ppm (Chapter 3), was analyzed after the addition of 200 ng of M e 3 S b C l 2 . The sample was analyzed in the same manner as the sample in Figure 2.15a and the result is shown in Figure 2.15c. The same demethylation pattern can be seen, indicating that this specific sample matrix causes the demethylation. The efficiency of the hydride generation reaction is decreased by the sample matrix, as shown by Figure 2.15d, which is the chromatogram that resulted when 200 ng of M e 3 S b C l 2 75 2.15a 8 e 3 100 90 80 70 H 60 60 40 30 20 2.15c w Time (min) 1 2 Time (min) c k 100 i 90 80 60 -| 60 40 H 30 20 10 2.15b w h-H—r> 4 -100 90 80-1 70 c 60 I " 30 20 10 0 2.15d Time (min) i 2 Time (min) Figure 2.15a. Chromatogram of an aqueous fungus extract analyzed at neutral pH (water only, unbuffered). Figure 2.15b. Chromatogram of the same aqueous fungus extract analyzed at pH 6.2, citrate buffer. Figure 2.15c Chromatogram of antimony-free aqueous fungus extract spiked with 200 ng Me3SbCl2, analyzed as in Figure 2.15a. Figure 2.15d. Chromatogram of 200 ng Me3SbCl2 in water analyzed as in Figure 2.15a. W = SbH3, X = MeSbH2, Y = Me2SbH, Z = Me3Sb. 76 dissolved in water was analyzed in the same manner as the sample in Figure 2.15a and Figure 2.15c. The amount of total hydrides in Figure 2.15d is much greater than that in Figure 2.15c. Figures 2.15(a-d) show the importance of matrix effects in this fungal extract on the hydride generation of Me 3 Sb . 2.2.3.1.4. Other studies to determine causes of demethylation Additional studies were carried out to qualitatively determine causes of demethylation, and are summarized in Table 2.6. Table 2.6. Qualitative tudies to determine causes of demethylation. See Figure 2.12 for locations of reactors A and B in Method 3. Reactor B contained 20 ml of I M H 2 S 0 4 and an amount of 100 ng (as Sb) of Me3SbCl2 was reacted in each experiment. I, II and H I are sequential in time. Expt . N o . Method Reactions Demethylation? 1 3 I. M e 3 S b C l 2 + N a B F L (reactor A ) EL Gaseous products from I bubbled through H 2 S 0 4 (reactor B ) no 2 2 I. M e 3 S b C l 2 + N a B F L H. (Products from I) + H 2 S 0 4 yes 3 3 I. 2 ml NaBFL, (2% w/v) + H 2 S 0 4 (reactor B ) H. M e 3 S b C l 2 + N a B F L (reactor A ) ITT. Gaseous products from II bubbled through liquid products from I (reactor B ) yes 4 3 I. M e 3 S b C l 2 + N a B F L (reactor A ) n. Gaseous products from I bubbled through H 2 S 0 4 (reactor B ) HI. (Liquid products from II in reactor B) + 2 ml N a B F L (2% w/v) yes 77 M e 3 S b C l 2 is stable in I M DC1 in D 2 0 by using N M R analysis , and therefore demethylation is not taking place prior to the hydride generation reaction. M e 3 S b was generated at neutral conditions and then bubbled through H 2 S 0 4 , and it was found to be stable to acid (Table 2.6, Experiment 1). However, when both borohydride and acid are present after trimethylstibine is generated, demethylation is observed (Table 2.6, Experiments 2 and 3). Trimethylstibine appears to dissolve in acid (or possibly become reoxidized to trimethylantimony oxide) and demethylates during the reaction with sodium borohydride (Table 2.6, Experiment 4). Therefore demethylation is taking place during the reaction, as a result of a product of the acid/borohydride reaction. As well, trimethylstibine, once formed, is unstable to a product of the acid/borohydride reaction. 2.2.3.1.5. Suggested reasons for demethylation "Molecular rearrangement" of methylarsines has been observed in the past 7 9 ' 8 0 ' 8 1, mostly in the form of demethylation, but also resulting in the formation of higher methylated compounds from lesser methylated ones. The previous studies do not give an explanation for the rearrangement, but rather link it to certain factors. For example, when 0.5 M H 2 S 0 4 was used in the derivatization reaction, demethylation of monomethyl- and dimethylarsines resulted 8 1. The present study (section 2.2.3.1.1) shows a similar trend, in that the use of acid during the hydride generation of M e 3 S b C l 2 results in demethylation products. However in another study the use of acid p H during the hydride generation reaction reduced the amount of demethylation7 9. In the same study 7 9 the use of a sodium borohydride pellet rather than aqueous solution and elimination of dissolved oxygen also reduced or eliminated the amount of demethylation, and this was attributed to an increase in the rate of the reduction reaction. In other words, these authors postulated that a faster rate of reaction results in less demethylation. 78 Mechanistic pathways for demethylation or rearrangement have not been postulated in the literature. The mechanism of the hydride generation reaction itself is not well known but studies with deuterium labeled sodium borohydride and arsenic compounds indicate that the hydrides result from direct FT transfer from the borohydride 8 2. The present study indicates that demethylation takes place in solution during the hydride generation reaction. It may then be postulated to take place at one or both of the following stages: during the actual reduction, or after Me 3 Sb is formed. To address the possibility of demethylation taking place during the reaction, the following reactions may be postulated to take place for the reduction of M e 3 S b C l 2 : M e 3 S b C l 2 + 2FT - » M e 3 S b H 2 + 2C1" (1) M e 3 S b H 2 Me 3 Sb + H 2 (2) Because the Sb-C bond is of similar strength to Sb-H but less than that of S b - C l 8 3 , 8 4 , methyl groups may be lost during these reactions. The intermediate compound M e 3 S b H 2 may be unstable to FT or to products of the reaction between acid and borohydride. Me 3 Sb , once formed, appears to be unstable to the products of the acid/borohydride reaction (section 2.2.3.1.4). The presence of borane, B H 3 or diborane, B2H<s, known to be products from the reaction of BFL," with H C 1 8 5 may be a key factor affecting the stability of Me 3 Sb. Adducts with B H 3 may form, such as M e 3 S b B H 3 which decomposes at room temperature86. The decomposition of M e 3 S b B H 3 may follow a similar pathway as that proposed for the reaction of Me4Pb and B 2 H 6 to M e 3 B and metallic Pb, which involves stepwise loss of methyl groups from the lead compound 8 7. M e 2 S b H B H 3 has been isolated and reacts further to form the stable compound M e 2 S b B H 2 8 8 . The relative instability of the Me-Sb bond in Me 3 Sb is 79 also seen by using mass spectrometry (see also Chapter 6) where the first methyl group is easily lost from Me3Sb. More experiments are necessary to deduce the mechanism of both the reduction reaction and the demethylation and rearrangement phenomena. Therefore, when analyzing a sample for methylated antimony compounds by the method of hydride generation, the reaction conditions should be carefully tested with standard compounds such as M e 3 S b C l 2 . 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F. , Eds.; Pergamon: Oxford, 1973; V o l . 1, pp 625. 85 Chapter 3 ARSENIC AND ANTIMONY IN MUSHROOMS 3.1. Introduction Mushrooms are fruiting bodies which participate in the reproductive cycle of a fungal organism. The fungus consists of filamentous cells (hyphae) that grow through a substrate to form a mass called the mycelium. Fungi that produce mushrooms belong to one of two divisions, the Basidiomycota and Ascomycota, based on the type of reproductive cell inside the mushroom used for spore production (basidium or asci). Spores are the reproductive cells that trigger the birth of new fungal organisms. Most of the commonly known fungi that produce mushrooms belong to the division Basidiomycota. Interest in the arsenic content in mushrooms has increased recently. Some species of mushrooms appear to accumulate arsenic and other metals from soil 1 , and their potential as biological pollution indicators has been discussed2. For mushrooms that accumulate arsenic, and are edible, such as Laccaria amethystina, toxicological consequences (if any) to consumers have been of concern 3. For these reasons, the uptake and speciation of arsenic in mushrooms has been studied. Determining the bioavailability and species of arsenic in mushrooms, especially those containing elevated levels, helps in toxicological risk assessment. Chemical processes taking place in the terrestrial environment have not been studied to the extent to which those in the marine environment have been. The recent findings of arsenobetaine and arsenocholine in mushrooms have led researchers to draw similarities between marine and terrestrial pathways for the formation of arsenic compounds 4 ' 5 ' 6. The presence of 86 arsenobetaine in mushrooms in higher taxonomic positions (i.e., being more highly evolved) 7 is similar to the presence of arsenobetaine in higher marine organisms, such as marine animals8. Although some authors believe that the fungi producing the mushrooms are responsible for the bio-synthesis of more complex arsenic forms, such as arsenobetaine7, no proof exists for this hypothesis. This theory is favoured for two reasons. One is that arsenobetaine has not been found in soil 4 . Arsenobetaine was, however, found in ant-hill material9, and its presence in estuarine waters was recently confirmed 1 0. The second reason is that similar chemical forms of arsenic have been seen in mushroom species collected from different locations 3. In support of the fungus biosynthesis theory, the fungi Agaricusplacomyces and Pleurotus sp. (producing edible mushrooms) methylate arsenic to a small extent1 1. Very little is known about the interaction of antimony with fungi, or with terrestrial organisms in general. The chemical similarities of antimony and arsenic has led to the hypothesis that fungi may interact with antimony in the way that they do with arsenic. For example, antimony was postulated to be methylated by the fungus Scopulariopsis brevicaulis, and this has, in fact, been shown to take place to a very small degree 1 2 ' 1 3. The identity of antimony species in fungi which produce mushrooms is completely unknown. The following chapter describes work exploring three objectives. The first is to determine the arsenic species in edible mushrooms that are readily available in supermarkets. The second is to determine i f mycelia in axenic culture (i.e., containing only one organism) are responsible for the formation of complex arsenicals by growing pure cultures of fungi that are capable of producing mushrooms in arsenic amended media. The third is to learn more about biological interactions with antimony, by growing fruiting bodies on antimony-containing substrate, and by culturing fungi that can produce mushrooms in antimony amended media. 87 3.2. Experimental 3.2.1. Chemicals and reagents Sodium arsenate, N a 2 H A s 0 4 . 7 H 2 0 (Aldrich), arsenic trioxide, A s 2 0 3 (Alfa), methanearsonic acid, C H 3 A s O ( O H ) 2 (Vineland Chemical), and cacodylic acid, ( C H 3 ) 2 A s O ( O H ) (Vineland Chemical) were dissolved in deionized water to make standard solutions. Extracts of kelp powder (Galloway's, Vancouver, B C ) and Nor i (Porphyra tenera) o f known arsenosugar content were obtained as described elsewhere1 4 and were used as laboratory standards in order to establish the retention times of arsenosugars. The identity and retention times of the arsenosugars were verified by comparison to pure arsenosugars generously donated by K . Francesconi and T. Kaise. Arsenobetaine 1 5, arsenocholine1 6, trimethylarsine oxide 1 7 , and tetramethylarsonium iodide 1 8 were had been synthesized previously according to standard methods. Identification of arsenicals in samples was made by comparison of retention times to those in standards. Antimony (V) and (III) standards were obtained as potassium hexahydroxyantimonate, K S b ( O H ) 6 (Aldrich), antimony trichloride, SbCl 3 (Aldrich), and potassium antimonyl tartrate, K 2 S b 2 ( C 4 0 6 H 2 ) 2 (Aldrich). M e 3 S b C l 2 was synthesized as described elsewhere1 9. Stock solutions were made by dissolving these compounds (except SbCl 3 ) in deionized water to 1000 or 100 ppm as Sb. SbCl 3 was dissolved in 6 M HC1 to 1000 ppm as Sb. Standard working solutions were made by diluting the stock solutions with deionized water as necessary. For hydride generation analysis, NaBFL, (reagent grade, Aldrich) was dissolved in deionized water fresh daily to give a concentration of 2% w/v. Ammonium citrate buffer at a concentration of 0.05 M and p H 6 (1 M ammonium hydroxide, Micro Select, Fluka, and 88 analytical reagent grade citric acid, B D H ) and 1 M HC1 (Environmental grade, Alfa Aesar) were used. Other chemicals for the preparation of mobile phases were analytical-reagent grade or higher in purity. They included methanol ( H P L C grade, Fisher), concentrated phosphoric acid (Aldrich), ammonium hydroxide ( I M , Fluka), pyridine (Fisher), formic acid (BDH) , potassium hydroxide ( K O H , Aldrich), tetraethylammonium hydroxide, ( T E A H , 20% in water, Aldrich), and malonic acid (BDH) . Mobile phases were filtered through 0.45 um cellulose nitrate filters (Millipore) prior to use. The arsenosugar stock solution used in the Scleroderma citrinum experiments was obtained by extracting commercially available macroalgae species (available for human consumption), which are known to contain arsenosugar X and X I , and D M A 1 4 . The sample consisting of 28 g Nova Scotian dulse (Palmariapalmatd), 6.5 g Taiwanese Nor i (Porphyra tenera) and 6.7 g Japanese Y a k i Nor i (Porphyra tenera, all purchased at Vancouver supermarkets) was extracted with approximately 1.5 L of MeOH/water (1:1) three times by sonication (2 hours), and the liquid was separated from the solid between sonications by filtration through Whatman #1 filter paper. The extracts were combined and concentrated to a final volume of approximately 400 mL by evaporation with a constant stream of air overnight at room temperature. 89 3.2.2. Apparatus and method of analysis 3.2.2.1. HG-GC-AAS analysis for antimony speciation The apparatus was composed of a semi-continuous flow, hydride generation system developed for arsenic analysis,2 0 coupled to an atomic absorption spectrometer (Varian AA1275) fitted with an Sb lamp (Varian) operating at a wavelength of 217.6 nm. One modification was made to the basic apparatus in the form of using a gas-liquid separator2 1 that resulted in less analyte carryover. The apparatus consisted of Tygon tubing for the peristaltic pump, and P T F E tubing (1/8" OD) for the remainder. The glass gas-liquid separator was silanized with ( C H 3 ) 2 S i C l 2 before use. Data were collected from the A A S and process directly by using an H P 3 3 90A integrator, or were analyzed with the aid of Shimadzu EZChrom software. A peristaltic pump was used to deliver standard or sample solution (usually 0.1 mL to 3 mL) to mix with the acid or buffer and then to mix with a solution of NaEfflU (2% w/v) in a reaction coil. The gases evolved were separated in the gas-liquid separator and then swept by a flow of helium into a P T F E U-tube, where they were trapped at -196 °C. Continuous hydride generation and trapping were carried out for 3 minutes. The peristaltic pump was then stopped (making the system semi-continuous) and the U-tube was heated to 70 °C, allowing the gases to be swept with He at a flow rate of 40 mL/min onto a Poropak PS column, which was then heated from 70 °C to 150 °C at a rate of 30 °C/min, whereby the gases were separated. They were then detected by A A S . 90 3.2.2.2. HPLC-ICP-MS analysis for antimony and arsenic speciation The H P L C apparatus consisted of a Waters 510 double piston pump, a Rheodyne six-port injection valve with a 20 u L loop, in line filters, a guard column for each analytical column packed with the same stationary phase, and the analytical column. Columns and mobile phases are listed in Table 3.2. In some cases of plasma instability and detector drift, 10 ppb Rh was added to the mobile phase to provide a constant background signal. A V G Plasmaquad PQ2 Turbo I C P - M S ( V G Elemental) was used as a detector. Parameters for the I C P - M S are given in Table 3.3. The m/z monitored were 121 and 123 (Sb), 75 (As), 77 82 (Se and Ar- 3 7 C1) and 103 (Rh) where applicable. The H P L C was coupled to the spray chamber of the I C P - M S by using a minimum of P T F E tubing (10 cm x 0.5 mm id . ) with the appropriate P T F E fittings. Extracts and media were diluted as necessary, filtered through 0.45 um syringe filters (Millipore) and analyzed by H P L C - I C P - M S using the conditions given in Table 3.1 (arsenic speciation), Table 3.2 (antimony speciation) and Table 3.3 ( ICP-MS detection). Data were processed and analyzed by using chomatographic software2 2, and when semi-quantitative concentrations of arsenic and antimony compounds were determined, external calibration curves were used. 3.2.2.3. ICP-MS analysis for total arsenic and antimony concentrations The I C P - M S described above, outfitted with a peristaltic pump and injection loop for flow injection introduction, was used for the determination of total arsenic and antimony in samples. The parameters listed in Table 3.3 were used, except that time resolved analysis was not used for these analyses. Solutions and standards were diluted with 1% (v/v) nitric acid (doubly distilled in quartz, Seastar) and Rh (10 ppb) was added as an internal standard. 91 Table 3.1. H P L C conditions for arsenic speciation Chromatography Column Mobile phase Flowrate (mL/min) Anion exchange Hamilton PRP-X100 , 150 x 4.6 or 250 x 4.6 mm 20 m M ammonium phosphate, p H 6.0 1.0 or 1.5 Cation exchange Supelcosil L C - S C X or Whatman S C X Partisil 5, 250 x 4.6 mm 20 m M pyridinium formate, p H 2.7 1.0 Ion-pairing G L Sciences O D S , 250 x 4.6 mm l O m M T E A H , 4.5 m M malonic acid, 0.1% M e O H , pH6 .8 0.8 Table 3.2. H P L C conditions for antimony speciation Chromatography Column Mobile phase Flowrate (mL/min) Anion exchange 2 3 Hamilton PRP-X100 , 150 x 4.6 mm 2 m M K O H 1 0 Ion-pairing Hamilton PRP-1 , 150 x 4.6 mm 10 m M T E A H , 4.5 m M malonic acid, 0.1% M e O H , p H 6.8 0.8 Table 3.3. Operation parameters for I C P - M S Feature Specific Conditions Forward radio-frequency power Reflected power Cooling gas flow rate (Ar) Intermediate (auxiliary) gas flow rate (Ar) Nebulizer gas flow rate (Ar) Nebulizer type Analysis mode Quadrupole pressure <10 W Time Resolved Analysis ( T R A ) for H P L C 9 x 10"7mbar de Galan 0.65 L/min 1350 W 13.8 L/min 1.002 L/min Expansion pressure 2.5 mbar 3.2.3. Cultivation of Pleurotus flabellatus fruiting bodies on a solid substrate To qualitatively determine antimony uptake by mushrooms, the fungus P. flabellatus (commonly known as the pink oyster mushroom) was grown on a solid soil medium (wood chips, sawdust and bran, about 25% moisture) contained in polyethylene bags. The bags containing 4 kg substrate were sterilized and inoculated with P. flabellatus mycelium, at Western Biologicals Ltd. , Aldergrove B C . The bags were opened in a biological safety cabinet at U B C and antimony solutions (10 mL of 500 ppm as Sb) were added through a 0.2 pm syringe filter (Nalgene), dropwise, and the top 15 cm of soil was mixed with a flame-sterilized spatula. The mixing was inadequate and hence concentrations were not homogeneous throughout the bag. Four bags were prepared, containing: (A) potassium antimonyl (III) tartrate; (B), potassium antimonate (V); (C) Me3SbCl2; and (D) no antimony. The bags were kept in a dark incubation chamber at 15 °C for 1 week, and then the temperature was increased to 20 °C for another week with illumination for 12 hours/day. The humidity was kept high by placing open dishes of water 93 around the bags, and by spraying the bags daily with deionized water. After 2 weeks fruiting bodies appeared and they were harvested and frozen until analysis. 3.2.4. Preparation of pure submerged cultures of fungi P. flabellatus was grown from a culture obtained from Western Biologicals Ltd. , Aldergrove, B C , in potato dextrose broth (Difco), a general liquid medium for fungus. The following procedures were used for all culture experiments. Solutions used to amend the media were sterilized by using syringe filters (0.22 um, Nalgene or Millipore, cellulose acetate), and they were added to give the appropriate starting concentration in each treatment (see Table 3.5). Seed culture containing mycelia that were free of antimony and arsenic was added in a volume that was approximately 10% of the volume of the broth (see Table 3.5 for volumes). The fungi were incubated on a shaker (forming spherical mycelia for P. flabellatus, S. citrinum and S. crispa, and broken filaments of mycelia f o r M procera) for the duration of the experiment (Table 3.5) at 26 °C. For P. flabellatus, the spherical mycelia were harvested by centrifugation of the biomass/liquid mixture and washing of the mycelia with deionized water. For the other cultures, the biomass was harvested by vacuum filtration (Whatman #1 filter paper) and rinsed a minimum of three times with deionized water. Scleroderma citrinum, commonly known as the earthball, and resembling puffballs in shape and reproduction, was obtained from an uncontaminated wood chip substrate in Vancouver. A number of fruiting body specimens were collected, cleaned manually by abrasion and with water to remove dirt and other debris, and sterilized on the outside surfaces with 30% hydrogen peroxide. Table 3.4. Summary of pure culture experiments. A B = arsenobetaine; AsS = arsenosugars from algae extract; control experiments contain non-living cells. Sb or A s species N o . of replicates P. flabellatus Sb (III) 2 Sb (V) 2 M e 3 S b C l 2 2 N o Sb or As 2 Sb (III) control 2 Sb (V) control 1 M e 3 S b C l 2 control 1 S. citrinum Sb (III) 2 Sb (V) 2 M e 3 S b C l 2 2 As (V) 2 A B 2 AsS 3 N o Sb or As 2 Sb (III) control 1 Sb (V) control 1 As (V) control 1 A B control 1 AsS control 1 M. procera As (V) 2 N o Sb or As 1 As (V) control 1 S. crispa As (V) 2 N o Sb or As 1 As (V) control 1 Duration of Approximate Culture experiment concentration volume (days) ( P P m ) (mL) 14 1 400 14 1 400 14 1 400 14 0 400 14 1 400 14 1 400 14 1 400 43 10 400 43 10 400 43 10 400 43 1 400 43 1 400 27 0.3 100 43 0 400 63 10 50 63 10 50 63 1 50 63 1 50 27 0.3 50 35 1 400 35 0 400 35 1 200 35 1 400 35 0 400 35 1 200 A l l other culturing steps were carried out aseptically in a biological safety cabinet. The specimens were cut open and a small piece of the fungus inside (containing spores) was used to inoculate sterile potato dextrose agar plates and potato dextrose broth. Within 2 weeks, growth was seen both on the plates (as a filamentous mycelium) and in the broth (as small white spherical mycelia, as for P. flabellatus). These cultures were then used to seed other stock cultures and the experimental cultures. M. procera and S. crispa were obtained from the American Culture Collection as axenic cultures, which were then used to inoculate Y M broth (see Table 3.4 for Y M broth ingredients, following the recipe recommended by Difco 2 4 ) . The submerged cultures obtained were used to seed the experimental cultures. Table 3.5. Y M Broth ingredients and composition Ingredients per liter Yeast extract, Difco Malt extract, Sigma Peptone, Difco Dextrose, Fisher Scientific p H + 0.2at25 °C Controls were prepared, containing Sb and As species, potato dextrose broth or Y M broth and mushroom mycelia that had been autoclaved. The controls were incubated for a minimum of the same time period as the live cell experiments, and the liquid was collected by filtration (Whatman #1 filter paper). N o organisms appeared to be growing in any flasks containing autoclaved cells, determined by visual inspection of the flasks, and by visual inspection of potato dextrose agar and nutrient agar that had been streaked with the autoclaved cells. Weight in grams 3 3 5 10 6.2 N o contamination by other organisms was observed, either macroscopically (i.e. no cloudy solutions, which indicate bacterial infection) or microscopically, except in an experiment containing Sparassis crispa and amended with As (V). Hence the experiments (except for the one containing live cells of S. crispa) were assumed to axenic, meaning that only one organism was present in each experiment. A portion (1-5 mL) of the medium for each experiment was collected after the addition of arsenic or antimony (referred to as "medium before") and reserved for analysis. The medium for each experiment was collected after harvesting of the biomass (referred to as "medium after") as well. A l l biomass and medium samples were frozen (-20° C) immediately until sample preparation and analysis. 3.2.5. Sample preparation and analysis Edible mushrooms were obtained canned or dried from Vancouver supermarkets. Canned mushrooms (oyster mushroom and straw mushroom) were homogenized in a blender with the liquid in the can, and freeze-dried. The freeze-dried material, and dried mushrooms were pulverized by using a mortar and pestle. The powders were then extracted. Extractions were carried out by weighing 2 g (± 0.5 mg) of the dried powders into 50 mL or 15 mL centrifuge tubes, adding 10-15 mL MeOH/Water (1:1), sonicating for 20 minutes, centrifuging for 20 minutes and decanting the liquid layer into a R B F . Each sample was sonicated and centrifuged a total of 5 times. The decanted extracts for each sample were pooled and rotovapped to near dryness (1-2 mL) and then dissolved in 5 or 10 m L of deionized water. P. flabellatus fruiting bodies were freeze-dried and pulverized by using a mortar and pestle. A soil sample consisting of equal amounts of soil from each bag was oven dried and weighed to calculate R (R = fresh weight/dry weight). Extractions were carried out as described 97 above, except that for mushrooms, 0.5 g (+ 0.5 mg) was weighed and extracted, and final extract volumes were 10 mL; and for soils, 3 g (fresh weight) was extracted (± 0.01 g) and final volumes were 10 mL. Samples of the fruiting bodies for P. flabellatus were digested with acid for determination of total antimony content. The freeze-dried mushroom powders were weighed (0.5 g ± 0.5 mg) into a 500 m L round bottomed flask (RBF). Concentrated nitric acid (3 mL, doubly distilled in quartz, Seastar, Sidney, B C ) , hydrogen peroxide (3 mL, 30% in water, reagent grade, Fisher) and concentrated sulfuric acid (1 mL, reagent grade, Fisher) were added to each sample. The samples in the R B F s were boiled for 3 hours by using a heating mantle and a reflux apparatus25. After all the samples had cooled, the clear solutions remaining were diluted to 10 mL with deionized water and stored at 4 °C until analysis. Medium samples were diluted with deionized water and filtered as necessary, and analyzed with no further preparation. Fresh weight biomass samples (P. flabellatus and <S". citrinum) were prepared by weighing 10 g (± 0.01 g) of filtered and washed mycelia spheres and homogenizing the spheres with an Ultraturrax T25 homogenizer (Jak & Kunkel). The samples were then centrifuged, and the resulting supernatant fraction was filtered. Dry weight samples (all species of fungi) were obtained by freeze-drying the mycelia and pulverizing them with a mortar and pestle. The samples were then prepared by weighing 0.25 g (± 0.5 mg) of the dry powder, extracting with 5 g (± 0.5 mg) deionized water by sonication for 2 hours, centrifuging the slurry, and collecting and filtering the resulting supernatant for analysis. 98 3.2.6. Isolation of an unknown compound containing antimony A n unknown antimony-containing compound was observed in all medium samples amended with inorganic antimony species when they were analyzed by using ion-pairing chromatography H P L C - I C P - M S . Due to the large proportions of this compound in most samples, an attempt was made to isolate and partially characterize it. A sample of potato dextrose broth medium containing Sb (V) in which P. flabellatus had been cultured was used. Undiluted medium (1 mL) was injected onto a PRP-1 column (4.6 cm x 15 cm) and the fraction eluting between 5 and 15 minutes was collected (a mobile phase of 10 m M T E A H / 4 . 5 m M malonic acid at p H 6.8 was used; see Table 3.2 for chromatographic details). This was repeated 9 times so that a total of 10 mL was injected onto the column. The fractions were pooled and rotovapped to dryness and then dissolved in about 2 m L with deionized water. Ethanol at -20 °C (4 mL) was added and the mixture was kept at -20 °C for 24 hours. The mixture was centrifuged, and the supernatant was then applied to a 30 mL Sephedex L H 2 0 (Pharmacia) in methanol column ( 3 x 1 0 cm). Methanol (90 mL) was applied to the column and collected, apart from the first 15 mL, and then evaporated to dryness. The solids remaining were dissolved in 1 m L of deionized water. 99 3.3. Results and Discussion 3.3.1 Arsenic species in edible mushrooms M u c h of the recent interest in arsenic in mushrooms stems from their potential as dietary sources of arsenic, especially in specimens collected from areas high in arsenic. For example, because of the presence of D M A in the choice edible Laccaria amethystina, it has been recommended that the ingestion of this mushroom grown on arsenic-contaminated soil be avoided 3. Therefore we were interested in analyzing some edible mushrooms commonly available in Vancouver supermarkets, to determine the identity of arsenic species present in the MeOH/water extractable portion. In past studies, the arsenic profile for the same species of mushroom collected from different locations and containing different levels of arsenic was found to be similar3. Thus we hypothesized that knowledge of the speciation of arsenic in mushrooms meant for human consumption might allow us to predict the speciation in the same mushrooms containing higher levels of arsenic. The mushrooms analyzed in this study were bought both in the dried form and in cans. Dried mushrooms included wooden ears (probably Auricula auricularia), which is a fungus used in many Chinese dishes; shiitake mushrooms (Lentinus edodes); two samples of porcini mushrooms, Porcini 1 (most likely Boletus sp.) and Porcini 2 (unidentified mushrooms that were most likely Agaricus sp., or portobello mushrooms); chanterelle mushrooms (Cantherellus cibarius); and a dried mushroom powder of unknown composition. Two species of mushrooms were obtained in cans; they were oyster mushrooms (most likely Pleurotus ostreatus) and straw mushrooms (probably Volvariella volvacea). The arsenic species extracted from these mushrooms by using MeOH/water (1:1) are summarized in Table 3.6. 100 Table 3.6. Arsenic species in edible mushrooms, in ppb dry weight (SD a ) . "Trace" amounts are greater than the limit o f detection (LOD) but less than 2 x L O D . Mushroom As (III) As (V) MMA DMA AB Me 4As + Sum of speciesb Wooden ears trace 17 (4) 23 (5) trace <3 <3 46 Shiitake 210 (30) 130 (10) trace 18(4) <5 <5 360 Porcini 1 56 (3) 30 (10) trace 46 (1)) <5 10(4) 150 Porcini 2 <5 30 (10) trace 70 (20) 54(6) <5 160 Chanterelle trace 22 <5 trace <5 <5 32 Mushroom powder <9 210 (50) trace 230 (60) 910 (50) <9 1360 Oyster mushrooms 30 (10) 28(1) 30 140 (10) <5 <5 230 Straw mushrooms <5 36 <5 trace <5 <5 41 a SD = standard deviation, obtained from analysis of extracts by using two different chromatographic systems (anion and cation exchange, see Table 3.1) with ICP-MS detection. b Calculation of sum of arsenic species included trace amounts estimated at the detection limit, which was 5 ppb for all mushrooms, except for Auricula sp. (3 ppb) and mushroom powder (9 ppb). The dried mushroom mentioned earlier, referred to as Porcini 2 and tentatively identified as Agaricus sp., was packaged under the common name "porcini" mushroom, the common name for Boletus sp. However, the appearance of the mushrooms revealed the fact that they had been identified incorrectly, since mushroom gills were observed, and Boletus sp. have pores rather than gills. Gills, pores or spines are found on the underside of many mushrooms with caps. The observation of a different profile of arsenic species in the unidentified mushrooms, compared with that found for a (presumed) correctly identified sample of porcini mushrooms (Porcini 1, Boletus sp.) supports the suggestion that the Porcini 2 mushrooms were identified incorrectly as porcini mushrooms. The most important difference in the arsenic profiles is the presence of arsenobetaine (34% of arsenic extracted) in the unidentified mushrooms. A number of mushrooms belonging to the genus Agaricus were analyzed in a previous study7 and all contained arsenobetaine, ranging in proportion from 55 to 96% of the arsenic extracted. Therefore it is not unreasonable to suggest that the unidentified mushroom may belong to the genus Agaricus. 101 The only other mushroom sample that contains arsenobetaine is the mushroom powder of unknown composition. The arsenic species found in the powder are similar to those found in Agaricus bisporus, both in identity and proportions 7 (Agaricus bisporus: A s (V) 12%, M M A 6%, D M A 27%, A B 55%; see Table 3.7). Agaricus bisporus is a mushroom commonly cultivated and eaten, and is also known as the button or white mushroom. Most likely the mushroom powder is composed of this species of mushroom. Table 3.7. Comparison of proportions of arsenic species in mushrooms (%) in the current study with those found in published studies (indicated by a footnote). Mushroom A s (III) As (V) M M A D M A A B M Porcini 2 0 19 3 44 34 __ Mushroom powder 0 15 1 17 67 0 Agaricus bisporus3 0 12 6 27 55 0 Oyster mushroom 13 12 13 61 0 0 Pleurotus ostreatush 60 39 1 0 0 0 Straw mushroom 0 88 0 12 0 0 Volvariella volvacea l a trace 4 8 78 10 0 Volvariella volvacea T trace 6 trace 94 trace 0 Porcini 1 37 20 3 31 0 7 Amanita caesara" 32 38 0 13 0 17 Agaricus campestef 0 0 trace trace 96 4 a Results from Slejkovec et al. (1997)7. b Results from Slejkovec et al. (1996)11 . In general, appreciable proportions of inorganic arsenic were extracted from most of the mushrooms. I f these mushrooms were collected from areas contaminated with arsenic and 102 similar arsenic species were present, the presence of higher levels of inorganic arsenic could be toxicologically important. Extractable arsenic species have been identified for the first time in wooden ears fungus (Auricula auricularia), shiitake mushrooms (Lentinus edodes), porcini mushrooms (Boletus sp.) and chanterelle mushrooms (Cantharellus cibarius). The low levels of arsenic in the sample of chanterelle mushrooms made the speciation of arsenic difficult. Me4As+, a compound not observed in any other mushrooms in this study, is observed in Porcini 1 (Boletus sp.). This compound has been observed in other mushroom species, including Amanita sp. and Agaricus sp. in previous studies (see Table 3.7)7. The low levels of arsenic in straw mushrooms made identification of arsenic species difficult. Arsenate appears to be the dominant species in this study, but D M A was found to be the major species in the presumed same mushroom in a previous study7. The species of arsenic in this mushroom differed between two different specimens in the previous study (see Table 3.7); one specimen contained inorganic arsenic, arsenobetaine and M M A , as well as D M A , and the other contained mostly D M A and a small amount of arsenate. Therefore the speciation of arsenic appears to be inconsistent for this mushroom species. Different arsenic profiles may arise from different microbial environments for the different specimens, i f uptake o f the arsenic compounds is taking place. Different metabolic pathways may be followed as well. When Pleurotus sp. were cultivated in soil amended with inorganic arsenic, only inorganic arsenic was observed in the mushroom, with 1% conversion to M M A (see Table 3 .7) n . In this study, D M A is the major species of arsenic extracted from oyster mushrooms (Pleurotus sp.), with inorganic arsenic and M M A present as well. This may indicate differences in the growing environment of the fungus, and that the arsenic species present in this mushroom are due to uptake from the environment rather than biosynthesis by the mushroom. Slejkovec et 103 al.n, however, suggested that the time scale for the study involving cultivation ofPleurotus sp. was not long enough for the mycelium or fruiting body to biosynthesize other arsenic species. This hypothesis may also account for the differences in arsenic species observed. 3.3.2. The interaction of arsenic species with pure submerged cultures of fungi 3.3.2.1. Culture experiments with Scleroderma citrinum As mentioned earlier, Scleroderma citrinum is commonly known as the earthball, and is similar to a puffball in appearance. It forms spherical fruiting bodies that can range up to 12 cm in diameter, and the spore mass inside is violet-black, becoming powdery and pale greenish or lilac gray when mature. Spores are propagated when the fruiting body is mature and opens. This mushroom was chosen because of its availability and also because of its resemblance to puffballs. Puffballs were found to contain arsenobetaine in previous studies7. Experiments were conducted in which the growth medium was amended with arsenate (As (V)), arsenobetaine ( A B ) and a mixture of arsenosugars (AsS). Arsenate is the most likely form of arsenic available in so i l 2 6 and probably to fungi in the environment. Arsenobetaine is a possible end product for some mushroom species and so it was of interest to determine i f any chemical changes take place as a result of mushroom metabolism. Arsenosugars are postulated to be intermediates in the biosynthesis of arsenobetaine27 and therefore their fate was of interest. 104 Concentrations of total arsenic in media and biomass for these experiments, as well as a bioconcentration factor (BCF) for the fresh weight biomass are summarized in Table 3.8. A B C F is defined as the quotient of the concentration of a material (in this case, arsenic) in an organism divided by the concentration of the material in the solution in which the organism has been l iving 2 8 . It can be calculated by using the following relation: B C F = ([As]biomass/[As]ma); ma = medium after. Total concentrations were obtained by using flow injection I C P - M S analysis as described in section 3.2.2.3, except for the samples from the arsenosugar amended experiment. These concentrations were obtained by summing the concentrations of arsenic species determined by H P L C - I C P - M S (as described in section 3.2.2.2). If a B C F is greater than 1, concentration of arsenic by the organism from the medium is taking place 2 9. Concentration by S. citrinum was taking place only for arsenobetaine because a B C F of 23 (Table 3.8) is observed. In fact, only 7% of the arsenic remains in solution, indicating that most of the arsenic is taken up by the organism. S. citrinum does not appear to accumulate arsenate or arsenosugars. The low levels of arsenic in the biomass for the arsenate-amended experiment may indicate that exclusion or fast excretion of inorganic arsenic is taking place by S. citrinum. 105 Table 3.8. Total concentrations of arsenic obtained by I C P - M S analysis (except where indicated) for experiments conducted with S. citrinum. A B = arsenobetaine, AsS = arsenosugar mixture Experiment/sample Concentration of A s (ppm) B C F ([As]bi 0ma S S/[As]m a) b (SD a ) As (V) amended Medium before 1.23 (0.03) Medium after 1.34(0.05) Biomass 0.20(0.03) 0.15 A B amended Medium before 1.30 (0.03) Medium after 0.09 (0.03) Biomass 2.04 (0.04) 23 AsS amended0 Medium before 0.37 c Medium after 0.30° Biomass 0.24° 0.80 a SD = standard deviation, calculated from duplicate biological experiments. b BCF = Bioconcentration factor, calculated as [As]biomass/[As]ma; where ma = medium after. c These total arsenic concentrations were not obtained by flow injection ICP-MS analysis, but rather from the sum of arsenic species determined by HPLC-ICP-MS. See text for more detail. The proportions of arsenic species in samples from arsenic-amended S. citrinum experiments are summarized in Table 3.9. The speciation of arsenic is expressed in proportions rather than absolute amounts because the analyses for most experiments were semi-quantitative. 106 Table 3.9. Proportions of arsenic species (%) in experiments conducted with S. citrinum; control experiments contain non-living cells and arsenic; A B = arsenobetaine, AsS = arsenosugar mixture Experiment/sample As(III) As(V) D M A Sugar Sugar A B — X XT As CV) amended Medium before 2.8 97.2 0 0 0 0 Medium after 80.3 19.7 0 0 0 0 Biomass 87.5 12.5 0 0 0 0 Control medium after 0.7 99.3 0 0 0 0 A B amended Medium before 0 1.0 0 0 0 99.0 Medium after 0 0 0 0 0 100 Biomass 0.6 0 0 0 0 99.4 Control medium before 0 0 0 0 0 100 Control medium after 0 0 0 0 0 100 AsS amended Medium before 0 0 10.2 14.1 75.7 0 Medium after 0 0 9.9 34.1 56.0 0 Biomass 0 0 10.0 34.2 55.9 0 Control medium before 0 4.6 9.3 12.8 73.3 0 Control medium after 0 0 8.9 14.5 76.6 0 For the experiments in which the growth medium was amended with arsenate, reduction to arsenite by the mycelia appears to be taking place (Table 3.9). N o reduction took place in the control containing arsenate and non-living cells, indicating that the reduction was due to the biological activity of the mycelia, rather than chemical transformations in the medium. The speciation of arsenic in the biomass was similar to that in the medium. Most likely the fungus takes up arsenate, reduces it to arsenite and excretes it as arsenite. A s shown by the concentrations of total arsenic in media and biomass for the experiments in which arsenobetaine was added (Table 3.8), accumulation of arsenobetaine from the medium 107 takes place, and Table 3.9 shows that arsenobetaine remains unchanged. This result may indicate that should arsenobetaine be present in the growing environment of a mushroom-producing fungus in the wild, it wi l l be taken up efficiently by the fungus and not be detectable in the soil. The proportions of arsenic species in the medium at the beginning of the experiment ("medium before") in Table 3.9 represents the composition of the arsenosugar mix used for the experiments (extract of dulse and Nor i , see section 3.2.1). The proportions of arsenic species are similar for the control media at the beginning and at the end of the experiment. This indicates that changes in proportions are caused by the biological action of the fungus. The proportion of D M A remains constant, but the amount of arsenosugar X increases, while that of arsenosugar X I decreases. The proportions in the biomass and in the medium at the end of the experiment are similar. However, no arsenobetaine or other arsenic species are observed from the interaction of the fungus with arsenosugars. In summary, these experiments show that accumulation of arsenobetaine by S. citrinum takes place. A s well, the mycelium form of this fungus in pure culture appears to be unable to biosynthesize methylarsenic species or other organoarsenic species from arsenate and arsenosugars. The fungus is responsible for the reduction of arsenate to arsenite, and the probable transformation of arsenosugar X I to arsenosugar X . 3.3.2.2. Culture experiments with Macrolepiota procera and Sparassis crispa Macrolepiota procera is commonly known as the parasol mushroom and it is a choice edible, growing up to 25 cm in diameter. It possesses a stem and cap with gills underneath, and the surface of the cap is scaly. Sparassis crispa is also a choice edible and its appearance is described as resembling that of a cauliflower, ranging from 10 to 60 cm 3 0 . Both these mushrooms were chosen for biological experiments because of the interesting arsenic speciation 108 results found for specimens collected from the wild. These published results from Slejkovec et al1 are summarized in Table 3.10. The major compound found in a wild specimen o f M procera was arsenobetaine and it seems reasonable that this fungus might be capable of bio synthesizing arsenobetaine. Arsenocholine, an arsenic compound found in minor amounts in the marine environment8, was found in large amounts in wild specimens of S. crispa, in addition to an appreciable amount of unknown arsenic species7. Table 3.10. Arsenic species (% of arsenic extracted) found in wild specimens o f M procera and S. crispa (from Slejkovec et al.1); A B = arsenobetaine, A C = arsenocholine Mushroom A B As(III), M M A , A C Unknowns 3 D M A , A s (V) M. procera 100 trace 0 0 S. crispa 1 31 3 (As(V)) 66 0 S. crispa 2 trace trace 45 55 a Unknown compound on cation exchange HPLC system. The present arsenic speciation results for experiments conducted with the two species of fungus, using arsenate to amend the growth medium, are summarized in Table 3.11. 109 Table 3.11. Concentrations of arsenic species in experiments conducted w i t h M procera and S. crispa in ppm (biomass is dry weight except where indicated); control experiments contain non-living cells and arsenic. Experiment/Sample As (III) As (V) D M A T M A O Sum of B C F C species M. procera Medium before <0.004 1 01 <0.004 <0.004 1 01 Medium after 0.062 0 67 <0.004 <0.004 0 73 Biomass (dry weight) 1.33 0 67 <0.02 <0.02 2 00 Biomass (fresh weight) 3 0.11 0 06 <0.002 <0.002 0 17 Control medium after <0.004 0 75 <0.004 <0.004 0 75 S. crispa Medium before <0.004 1 08 <0.004 <0.004 1 08 Medium after <0.004 0 85 0.043 0.26 1 15 Biomass (dry weight) 2.56 2 62 0.29 1.8 7 27 Biomass (fresh weight) b 0.14 0 14 0.02 0.1 0 40 Control medium before <0.004 1 05 <0.004 <0.004 1 05 Control medium after <0.004 0 95 <0.004 <0.004 0 95 a Fresh weight concentration is calculated as [biomass concentration (dry weight)]/R, where R = fresh weight of biomass/dry weight of biomass; R f o r M procera is 11.9. b Fresh weight concentration is determined as forM procera, R for S. crispa is 18.2. 0 BCF = bioconcentration factor; see Table 3.8 for calculation. Because the B C F values are less than 1 (Table 3.11), no accumulation of arsenate is taking place by these fungi. The cultures of S. crispa were not axenic and therefore the conclusion cannot be drawn that the methylated compounds ( D M A and T M A O ) are present due to metabolism by the fungus, rather than by other organisms (e.g., bacteria) present. M. procera biomass contains arsenic mostly as arsenite but excretes very little of it, which may indicate that arsenite is sequestered by the organism after it is formed, or that the excretion process is slower 110 than the time scale of these experiments. Neither arsenobetaine nor arsenocholine are present in the biomass or in the media, which may reflect the inability of these species to biosynthesize these compounds. 3.3.2.3. Summary of the interaction of arsenic with fungi that can produce mushrooms Mycelia of mushroom-producing fungi grown in pure culture appear to be unable to synthesize arsenobetaine or arsenocholine, and reduction of arsenate to arsenite appears to be the only chemical process taking place. Arsenobetaine is efficiently taken up by S. citrinum which suggests that i f any arsenobetaine is present in the growing environment of wild fungi, it may be accumulated by the mushroom. Similar results were observed by Slejkovec et al.11. That is, only a small amount of methylation by Pleurotus sp. fruiting bodies and Agaricusplacomyces mycelia in pure culture was observed. Additionally, when the growth medium for pure cultures of Agaricus placomyces was amended with arsenobetaine and tetramethylarsonium ion, these two compounds were taken up to a high extent1 1. The authors suggested that the biosynthesis of arsenobetaine may not be observed because of the short time scale of the laboratory experiments1 1, which is probably the case for S. citrinum, M. procera and S. crispa as well. I l l 3.3.3. The interaction of antimony species with fungi 3.3.3.1. Cultivation of Pleurotus flabellatus fruiting bodies Although antimony has been listed as a U S - E P A priority pollutant, very little is known about its interaction with living organisms. Mushrooms were grown in soil that had been amended with antimony in experiments designed to contribute information about the interaction of antimony with fungi. It was of interest to determine, qualitatively, i f antimony is taken up under these conditions and to obtain speciation information. The strawberry oyster mushroom Pleurotus flabellatus was chosen because of its availability as a pure culture (Western Biologicals Ltd . , Aldergrove, B C ) as well as its fast growth rate. As mentioned in section 3.2.3, the fruiting bodies were cultivated on a solid substrate contained in polyethylene bags, a common method used by home mushroom growers. When the conditions are ideal, the fruiting bodies push through holes in the polyethylene to appear on the outside of the bag. Oyster mushrooms have short lateral stems and they are gilled on the underside of the cap. Pleurotus flabellatus is pink in colour and produces fruit in 2 weeks from the time of substrate inoculation. Four bags were prepared, containing: (A) potassium antimonyl (III) tartrate; (B), potassium antimonate (V); (C) M e 3 S b C l 2 ; and (D) no antimony. The concentrations of total antimony in mushrooms from bags A , B and C were elevated compared to D, the mushroom that had been exposed to substrate not amended with antimony (see Table 3.12). Prior to acid digestion, Sb(OH) 6", SbCl 4", K 2 Sb2 (C40 6 H 2 ) 2 and M e 3 S b C l 2 (0.125 pg each, to give a total of 0.5 ug of Sb) were spiked into the round bottom flask (RBF) containing the mushroom D, and an average recovery of 95% was obtained (see Table 3.12). These results indicate that the acid digestion procedure was adequate to dissolve antimony in these forms and probably that bound to the mushroom matrix. 112 Table 3.12. Antimony in mushrooms after acid A A S . digestion, analyzed by hydride generation-GC-Experiment Sb species3 [Sb] (ppm dry weight) in mushrooms A Sb (III) 6.5 A (following extractionb) Sb (III) 5.1 B S b ( V ) 1.6 C Me3SbCI2 1.0 C (following extraction) Me3SbC12 0.86 D noSb 0.04 D (spiked, replicate 1) no Sb 0.42 c D (spiked, replicate 2) no Sb 0.53 c Sb species in this column are those added to amend the mushroom growing soil. ' Extraction was carried out by using Concentration of spike = 0.50 ppm b xtractio  as carrie  o t y si  MeOH/water (1:1), as detailed in section 3.2.5. Very little antimony in the mushrooms is in a form extractable by M e O H / H 2 0 (1:1) for mushrooms A and C, leaving the remainder bound up with the residue (Table 3.12 and Table 3.13). Semi-quantitative amounts of antimony species found in mushroom and soil extracts are summarized in Table 3.13. Sb (III) is oxidized to Sb (V), but the possibility of oxidation by the mushrooms is unverified since no biomass-free control experiment (soil, bag and Sb (III) only) was carried out; thus, the oxidation of Sb (III) by the soil or by the air cannot ruled out. The oxidation state of antimony remains the same (i.e., Sb (V)) in both the soil and mushroom for experiment B . In experiment C, M e 3 S b C l 2 is taken up by the mushroom unchanged (i.e., no methyl groups were lost) and it also remains unchanged in the soil. Recoveries of soil spiked with all three antimony species, followed by extraction, are very low (<20%), indicating that the antimony species are strongly absorbed or adsorbed to the soil. 113 Table 3.13. Antimony extracted from mushrooms and soils 3 (ppm dry weight), H G - G C - A A S analysis. "Trace" amounts are greater than the limit o f detection ( L O D ) but less than 3 x L O D . When summing species, trace amounts were given a value of the L O D . Sample Sb (III) Sb ( V ) b M e 3 S b C l 2 Sum of Sb species Mushroom A <0.02 0.087 <0.02 0.087 Soil A <0.003 1.0 <0.003 1.0 Mushroom B <0.02 0.062 <0.02 0.062 S o i l B <0.003 0.46 <0.003 0.046 Mushroom C <0.0002 0.001 0.0033 0.0043 S o i l C <0.001 0.016 2.3 2.32 Mushroom D <0.02 trace <0.02 0.01 S o i l D <0.003 trace <0.003 0.003 Spiked soil D 0.016 0.29 0.25 0.556 % recovery 0 0.1% 16% 14% a Dry weight concentrations for soils were obtained by multiplying fresh weight concentrations by R = 1.85. b Limit of detection (LOD) for Sb (V) was 0.01 ppm for mushrooms, except for mushroom C, for which a LOD of 0.0002 ppm was estimated, and 0.003 ppm for soils. 0 Concentration of spike =1.84 ppm each 3.3.3.2. Culture experiments with Pleurotus flabellatus 3.3.3.2.1. ICP-MS and HG-GC-AAS analysis of biomass extracts and media More experiments with P. flabellatus were carried out, with the following points in mind. The problems of antimony absorption or adsorption to soil was avoided by growing the fungus in submerged culture to form mycelial spheres. We wished to confirm the oxidation of Sb (III) to Sb (V) by comparison with controls containing antimony and non-living cells. Finally, higher concentrations of antimony were used to allow the possible observation of new antimony species by H P L C - I C P - M S . A summary of concentrations in media at the beginning and the end of the experiments is given in Table 3.14. Two analysis techniques were used: hydride generation-GC-AAS, to obtain 114 speciation information; and I C P - M S , to determine the total amount of Sb. The concentrations of antimony acquired by using H G - G C - A A S differed from those obtained by using I C P - M S for the experiments in which inorganic antimony was used (Sb (III) and Sb (V)). This may indicate the presence of an antimony complex or compound that cannot be derivatized to a hydride under the analysis conditions. The concentrations of inorganic antimony species were determined either by using the method of standard additions, or by using an external calibration curve based on standards in a matching matrix. Both these methods of quantification minimize the likelihood of inhibition of hydride formation from inorganic antimony because of a matrix effect. N o such difference is seen for the experiment in which MesSbCU is used, except for the media at the end of the experiment, which show a decrease in the amount of hydride formed from M e 3 S b C l 2 . N o appreciable differences are seen in the concentrations of total Sb (determined by using ICP-MS) in the media at the beginning and the end of the experiment. 115 Table 3.14. Antimony in media and biomass extracts of Pleurotus flabellatus grown in submerged culture (ppm in solution, ppm fresh weight for biomass) (SD a ) . H G - G C - A A S analysis was used for speciation, I C P - M S was used for total Sb; control experiments contain non-living cells; na = not analyzed. Experiment/sample Sb (III) Sb (V) M e 3 S b - b Total (ICP-M S ) Sb am amended Medium before 0.40 (0.01) na <0.05 1.2 (0.2) Medium after 0.001 0.35 (0.04) < 0.001 1.09 (0.04) Biomass extract 0.004 0.47 < 0.002 na Control medium before 0.39 (0.08) na <0.05 1.14(0.01) Control medium after 0.37 (0.06) na <0.05 1.108 (0.003) Sb (VI amended Medium before na 0.4 (0.1) <0.03 1.2 (0.2) Medium after na 0.4 (0.3) <0.05 1.2 (0.2) Biomass extract < 0.002 0.57 < 0.002 na Control medium before na na na 1.27 Control medium after na na na 1.36 Me 3 SbCl? amended Medium before <0.05 na 1.11 (0.05) 1.2 (0.1) Medium after <0.01 na 0.45 (0.03) 1.1(0.1) Biomass extract < 0.0008 < 0.0008 0.42 na Control medium before <0.05 na 1.29 1.1 Control medium after <0.05 na 1.22 1.0 a SD = standard deviation, calculated for replicate experiments. b Me3Sb- is a compound containing Me3Sb, determined by HG-GC-AAS, meaning that the exact structure of the species prior to derivatization is unknown. The concentrations of antimony in the fresh biomass after homogenization were determined to be about 0.5 ppm fresh weight, by H G - G C - A A S , for all three treatments (Table 116 3.14). This concentration is similar to the concentrations obtained by H G - G C - A A S for the media samples at the end of the experiment (0.35 to 0.45, see Table 3.14), which may indicate that no bioconcentration of antimony is taking place by the mushroom mycelia. The antimony species found by using the method of H G - G C - A A S in the mushrooms treated with Sb (III) and Sb (V) is Sb (V) . The species of antimony found in the mushrooms treated with M e 3 S b C l 2 is a compound containing Me 3 Sb- . 3.3.3.2.2. HPLC-ICP-MS analysis of biomass extracts and media Two H P L C systems were used with I C P - M S as a detector: anion exchange chromatography with a mobile phase of 2 m M K O H 2 3 (Method A ) , and ion-pairing chromatography, with a mobile phase of 10 m M TEAH/4 .5 m M malonic acid at p H 6.8 and a polymeric reversed phase column (Method B ) ; see Table 3.3 for details. The separation of M e 3 S b C l 2 and Sb (V) (as Sb(OH)6~) by using these two chromatographic systems with I C P - M S detection is presented in Figure 3.1. Sb (III) standards are not eluted regardless of the method used. In Figure 3.1a, Me3SbCl2 elutes unretained on the column in the anion exchange system (Method A ) and Sb(OH) 6" is retained, affording a separation between the two compounds; this was observed previously as we l l 2 3 . When using the ion-pairing chromatography system (Method B) , both Sb(OH) 6" and M e 3 S b C l 2 are retained and separated (Figure 3.1b). 117 Figure 3.1. Chromatograms of standard antimony compounds (100 ppb each) on two H P L C - I C P - M S systems. 3.1a. 2 m M K O H , PRP-X100 antion exchange column. 3.1b. l O m M T E A H , 4.5 m M malonic acid, p H 6.8, 0.1% M e O H , PRP-1 reversed phase column. 118 When Method A was used for H P L C analysis, several unknown antimony-containing compounds, labeled A l , A 2 and A 3 , were observed. A chromatogram is shown in Figure 3.2 for a sample (medium after for the experiment amended with Sb (V)) in which the unknown compounds A l , A 2 and A3 are present. The relative amounts of antimony compounds when some of the samples were analyzed are summarized in Table 3.15. The H P L C analysis was qualitative only, hence absolute amounts are not given, and the assumption was made that the ionization in the plasma and response was similar for all antimony compounds. Table 3.15. Relative amounts of antimony compounds (%) in some P. flabellatus samples analyzed by using Method A . Control experiments contain non-living cells and antimony; A l -A3= unknown compounds. Experiment/sample M e 3 S b - Sb(OH) 6" A l A 2 A3 SUV) amended Biomass extract 0 57 43 0 0 Medium before 0 66 18 14 0 Medium after 0 46 15 16 23 Control medium after 0 44 25 31 0 Sbdin amended Biomass extract 0 56 44 0 0 Medium after 0 73 0 11 15 Me 3 SbCl? amended Biomass extract 95 5 0 0 0 Medium after 99 1 0 0 0 119 7e+3 120 240 360 480 time (s) 600 720 Figure 3.2. Chromatogram (Method A ) of medium after 14 days of growth for P. flabellatus amended with Sb (V). Upon examining Table 3.15, it can be observed that a large amount of A l (43 and 44%) is present in biomass extracts for the experiments treated with inorganic antimony. A3 is seen only in media sampled at the end of the experiments amended with Sb (III) and Sb (V). For the experiment amended with Sb (V), A3 is not present in the medium at the beginning of the experiment (medium before), nor in the control medium at the end of the growing period (control medium after). Thus these results suggest that a metabolite is formed from inorganic antimony starting compounds by P. flabellatus, and excreted into the medium. This metabolite might contain antimony, or it may bind to antimony already present in the medium. N o new species are formed when M e 3 S b C l 2 is used as the starting compound. When Method B was used for FIPLC analysis, unknown antimony-containing compounds, labeled B l , B 2 , B3 and B4 , were observed as well. Three chromatograms obtained by using Method B are shown in Figure 3.3. The first chromatogram (Figure 3.3a) is for the medium at the beginning of the experiment amended with Sb (V) (medium before), the second chromatogram (Figure 3.3b) was obtained from the medium at the end of the same experiment (medium after) and the third chromatogram (Figure 3.3 c) represents a fresh weight extract of the biomass collected from this experiment. Unknown B 4 is a major antimony-containing compound in the medium samples, but it is not present in the biomass extract. Chromatograms for samples from the experiment in which Sb (III) was used to amend the medium are similar to those in Figure 3.3 and are hence not shown. Due to its presence in all medium samples, including all controls, B 4 cannot be considered a metabolite. B3 is found in small amounts in biomass extracts for experiments amended with inorganic antimony but not in medium samples; however, its presence as a metabolite was not determined. 121 5e+4 4e+4 -3e+4 -2e+4 1e+4 H Oe+0 1e+5 c CD J3 < 8e+4 H 6e+4 4e+4 -2e+4 -Oe+0 -2e+4 2e+4 H 1e+4 8e+3 -4e+3 -Oe+0 0 Sb(OH) 6 Sb(OH) Sb(OH) 3.3a 3.3b 3.3c B2 B3 120 240 I I 360 480 time (s) B4 B4 600 720 Figure 3.3. Chromatograms of P. flabellatus media and biomass extracts (Method B ) for experiments amended with Sb (V). 3.3a. Medium before. 3.3b. Medium after. 3.3c. biomass extract (dry weight). B l , B2 , B3 and B4 are unknown Sb-containing compounds. 122 Because of the ubiquitous presence of B4 , an attempt was made to characterize it. As detailed in section 3.2.6, B 4 was isolated from a medium sample ("medium after", amended with Sb (V)) by using H P L C . Ethanol (-20° C) was added to the isolation product to precipitate proteins according to the method published by M a et a/ . 3 1 The sample was then applied to a Sephedex L H 2 0 column to remove excessive salts. The final isolated compound was analyzed qualitatively. H G - G C - A A S analysis when using I M HC1 (acid pH) to adjust the p H of the reaction afforded only SbH 3 . When H G - G C - A A S analysis was carried out at neutral pH, stibines were not produced. Hence the final isolated compound possesses the oxidation state of +5 and does not contain methyl groups bound to the antimony. However, the presence of antimony-containing compounds of unknown oxidation state that are not derivatized to hydrides cannot be discounted. This type of compound was suggested to be present in media from these experiments in section 3.3.3.2.1 and Table 3.14. The chromatogram obtained when the isolated unknown compound was qualitatively analyzed by using Method B is shown in Figure 3.4. The first chromatogram (Figure 3.4a) was obtained when the sample used for the isolation procedure was analyzed; this was the medium at the end of the experiment amended with Sb (V) (medium after), and prior to the isolation procedure. The second chromatogram (Figure 3.4b) represents the final isolated product, after ethanol precipitation and Sephedex L H 2 0 clean-up. Each of the chromatograms also shows the relative amounts of the compound (assuming similar detector response), and it can be seen that Sb(OH)6~ and B 4 were present in each sample. Only B 4 was the starting compound for the chromatogram in Figure 3.4b, indicating that B4 may in fact be an Sb (V) compound in equilibrium with Sb(OH) 6". 123 1.2e+5 I 8.0e+4 E 5 4.0e+4 H < O.Oe+Ci Sb(OH)6- 3.4a *l (82%) B4 (18%) I I I I I ^ I 12fJ '240 360 480 6CKk 720 1.6e+5 | 1.26+5 H £ 8.0e+4 CO •*—• In < 4.0e+4 0.0e+0 \Sb(OH)6" 3.4b \ (85%) B4 —.. - ,w-(15%) 120 240 r 360 Time (s) 480 600 720 Figure 3.4. Chromatograms of unknown B4 (Method B ) showing proportions of each compound. 3.4a. Medium after for P. flabellatus amended with Sb (V). 3.4b. B4 after H P L C fraction collection, ethanol precipitation and Sephedex L H 2 0 clean-up. 124 The equilibrium may be represented by the following equation: B 4 S b ( O H V When B 4 is isolated, the equilibrium is re-established so that Sb(OH) 6" is present. Other attempts to characterize this compound were unsuccessful. Enhanced oxidation of Sb (III) to Sb (V) appears to be taking place due to the presence of P. flabellatus, as suggested by the results in Table 3.14. Equal amounts of Sb (III) were found in the medium at the start of the experiment (amended with Sb (III)), and in the controls treated with Sb (III) at the beginning and the end of the experiment. Raw area counts (quantification was not carried out) are shown for Sb(OH) 6" present in media (Figure 3.5). Although some oxidation is taking place in the medium in the absence of living cells, enhanced oxidation is taking place by P. flabellatus (Figure 3.5). 3 E + 7 T 2 E + 7 - -1 E + 7 -O E + 0 -I mmmmmm 1 mmmmm 1 mrnt^im 1 m e d i u m b e f o r e m e d i u m a f t e r c o n t r o l m e d i u m a f t e r Figure 3.5. Raw area counts for Sb(OH) 6" in media for P. flabellatus grown in Sb (Ill)-amended culture. 125 CO X ) l_ CO 3.3.3.3. Culture experiments with Scleroderma citrinum The earthball, Scleroderma citrinum, was grown in pure submerged culture amended with antimony compounds to determine the interaction of antimony with this organism. The concentrations of total antimony in medium and biomass samples for cultures amended with antimony compounds are summarized in Table 3.16. The concentrations in media do not differ appreciably from the beginning (medium before) to the end (medium after) of the experiments. Bioconcentration factors are similar for the three antimony compounds studied and are approximately 0.2, indicating that no accumulation of antimony takes place by the fungus. Table 3.16. Concentration of total antimony (ppm) (SD a ) in media and biomass extracts of Scleroderma citrinum grown in submerged culture (ppm for solutions, ppm fresh weight for biomass). I C P - M S analysis was used for the analysis. Experiment/sample [Sb] B C F .([?^ ].^ .9m?.?^ I§.!?.]fn?) Sb (III) amended Medium before Medium after Biomass extract 12.0 (0.5) 10.9 (0.3) 2.0 (0.7) 0.18 Sb(V) amended Medium before Medium after Biomass extract 12.7 (0.7) 12.2 (0.3) 2.5 (0.3) 0.20 Me3_SbClz amended Medium before Medium after Biomass extract 10.9 (0.7) 10.7 (0.4) 1.8 (0.4) 0.17 a SD = standard deviation, calculated for replicate experiments. b BCF = Bioconcentration factor, calculated as [Sb]bi0mass/[Sb]ma; where ma = medium after. 126 The chromatograms resulting from the analysis of biomass extracts for the three experiments by using Method A (anion exchange H P L C - I C P - M S with 2 m M K O H ) are shown in Figure 3.6. The first two chromatograms were obtained from the analysis of the biomass extracts for S. citrinum grown with Sb (III) (Figure 3.6a) and Sb (V) (Figure 3.6b). A second antimony-containing peak of unknown identity and with a retention time similar to that of A l in Table 3.15, is observed in both these chromatograms. The third chromatogram (obtained by analyzing the biomass extract from the M e 3 S b C l 2 amended culture) shows the presence of M e 3 S b C l 2 (or the hydrolyzed form, Me 3 Sb(OH) 2 ) , most likely unchanged, in the biomass (Figure 3.6c). The medium samples analyzed by using Method A (not shown) contain only Sb (V) for the experiments amended with inorganic antimony, and only M e 3 S b C l 2 for the experiment amended with M e 3 S b C l 2 . N o new antimony compounds were detected by using this method of analysis, contrasting with the finding of a metabolite formed by P. flabellatus. 127 2e+4 Oe+0 8e+4 6e+4 -4e+4 -2e+4 Oe+0 Me3SbCI2 3.6c 120 r~ 240 time (s) 360 480 Figure 3.6. Chromatograms for biomass extracts (fresh weight) for S. citrinum experiments (Method A) . 3.6a. Experiment amended with Sb (III). 3.6b. Experiment amended with Sb (V) . 3.6c. Experiment amended with Me3SbCl2. A l is an unknown Sb-containing compound. 128 When the samples were analyzed by using Method B (ion-pairing H P L C - I C P - M S with 10 m M TEAH/4 .5 m M malonic acid, p H 6.8), the unknown described in the previous section, B4, was present in all samples, including biomass extracts. The identification of B 4 in these samples was based on the similar retention time of the peak compared with that observed in the samples for P. flabellatus. Considering only the area counts for Sb(OH) 6", increased oxidation of Sb (III) to Sb (V) in the presence of the live fungus does not take place in these experiments. 3.3.3.4. Summary of the interaction of antimony with fungi that can produce mushrooms Elevated levels of antimony were observed in fruiting bodies of P. flabellatus grown on substrate amended with antimony compounds, compared with levels found in the mushrooms grown on non-Sb containing substrate. Sb (V) was the only antimony species extracted from the mushrooms grown with both Sb (III) and Sb (V), and a Me 3Sb-containing compound was extracted from mushrooms grown with Me3SbCl2. N o appreciable differences were observed in total antimony concentrations in culture media for both P. flabellatus and S. citrinum, when the media were amended with antimony compounds. Thus the mycelia do not accumulate these antimony compounds. Previous studies with other fungi have shown different results. Scopulariopsis brevicaulis appears to take up Sb (III) from solution (losses of 30% were observed) 1 3, and Saccharomyces cerevisiae completely took up Sb (III) but not Sb ( V ) 3 2 . When Method A was used for the analysis of biomass extracts and media, a metabolite was detected, presumably produced from the interaction of P. flabellatus with inorganic antimony compounds. N o metabolites were detected for S. citrinum by using Method A . The antimony species in biomass extracts were similar for P. flabellatus and S. citrinum experiments, 129 showing a second antimony-containing compound eluting closely with Sb(OH) 6". Further studies to identify unknowns A l and A 3 are recommended. When Method B was used for analysis, the unknown compound B 4 appeared in nearly all biomass extracts and media. Thus an attempt was made to characterize it. B 4 may be in the +5 oxidation state and it appears to be in equilibrium with Sb(OH) 6". We suggest that further studies address either this compound's identification or elimination (e.g., by using a growth medium with minimum carbon sources and salts). Enhanced oxidation of Sb (III) to Sb (V) by P. flabellatus takes place, although oxidation by other factors is appreciable. 130 References 1. Slekovec, M . ; Irgolic, K . J. Chem. Spec. Bioavail. 1996, 8, 67-73. 2. Byrne, A R . ; Tusek-Znidaric, M . Appl. Organomet. Chem. 1990, 4, 43-48. 3. Larsen, E . H . ; Hansen, M . ; Goessler, W. Appl. Organomet. Chem. 1998, 72, 285-291. 4. Byrne, A . R.; Slejkovec, Z . ; Stijve, T.; Fay, L . ; Goessler, W. ; Gailer, J.; Irgolic, K . J. Appl. Organomet. Chem. 1995, 9, 305-313. 5. Kuehnelt, D . ; Goessler, W. ; Irgolic, K . J. Appl. Organomet. Chem. 1997,11, 289-296. 6. Kuehnelt, D . ; Goessler, W. ; Irgolic, K . J . Appl. Organomet. Chem. 1997, 11, 459-470. 7. Slejkovec, Z . ; Byrne, A . R.; Stijve, T.; Goessler, W. ; Irgolic, K . J. Appl. Organomet. Chem. 1997, 11, 673-682. 8. Edmonds, J. S.; Francesconi, K . A . Mar. Poll. Bull. 1993, 26, 665-674. 9. Kuehnelt, D . ; Goessler, W. ; Schlagenhaufen, C ; Irgolic, K . J. Appl. Organomet. Chem. 1997, 11, 859-867. 10. Florencio, M . H . ; Duarte, M . F.; Facchetti, S.; Gomes, M . L . ; Goessler, W. ; Irgolic, K . J.; van't Klooster, H . A . ; Montanarella, L . ; Ritsema, R.; Vilas-Boas, L . F.; de Bettencourt, A . M . M . Analusius 1997, 25, 226-229. 11. Slejkovec, Z . ; Byrne, A . R.; Goessler, W. Kuehnelt, D . Irgolic, K . J.; Pohleven, F. Acta Chim. Slov. 1996, 43, 269-283. 12. Jenkins, R. O.; Craig, P. J.; Goessler, W. ; Miller, D . Ostah, N . ; Irgolic, K . J. Environ. Sci. Tech. 1998, 32, 882-885. 13. Andrewes, P.; Cullen, W. R.; Feldmann, J.; Koch, I.; Polishchuk, E . Appl. Organomet. Chem. 1998, in press. 14. La i , V . W . - M . ; Cullen, W. R.; Harrington, C. F.; Reimer, K . J. Appl. Organomet. Chem. 1997, 77, 797-803. 15. Edmonds, J. S.; Francesconi, K . A . ; Cannon, J. R.; Raston, C. L . ; Skelton, B . W. ; White, A . H . Tetrahedron Lett. 1977, 18, 1543-1546. 16. Irgolic, K . J.; Junk, T.; Kos, C ; McShane, W. S.; Pappalardo, G . C. Appl. Organomet. Chem. 1987, 1, 403-412. 131 17. Nelson, J. C. Ph.D. Thesis, University of British Columbia, 1993. 18. Cullen, W . R.; Dodd, M . Appl. Organomet. Chem., 1989, 3, 401. 19. Morgan, G . T.; Davies, G. R. Proc. Royal Soc, Ser.A 1926, 523. 20. Cullen, W. R.; L i , H . ; Hewitt, G . ; Reimer, K . J.; Zalunardo, N . Appl. Organomet. Chem., 1994, 8, 303. 21. Le , X . C ; Cullen, W. R.; Reimer, K . J. Appl. Organomet. Chem. 1992, 6, 161. 22. Koelbl, G . ; Kalcher, K . ; Irgolic, K . J. J. Automat. Chem. 1993, 15, 37. 23. Lintschinger, J.; Koch, I.; Serves, S.; Feldmann, J.; Cullen, W . R. Fresenius J. Anal. Chem. 1997, 359, 484-491. 24. Difco Manual; Difco Laboratories: Detroit, 1984; pp 1131-1132. 25. Bajo, S.; Suter, U . ; Aeschliman, B . Analytica ChimicaActa 1983,149, 321-355. 26. Helgesen, H ; Larsen, E . H . Analyst, 1998, 123, 791-796. 27. Edmonds, J. S.; Francesconi, K . A . Appl. Organomet. Chem. 1988, 2, 297-302. 28. Aquatic Pollution, 2nd ed.; Laws, E . A , Ed . ; John Wiley & Sons: New York, 1993; ppl97-198. 29. Dushenko, W. T.; Bright, D . A ; Reimer, K . J. Aquat. Bot. 1995, 50, 141-158. 30. Pacioni, G . Simon & Schuster's Guide to Mushrooms; Lincoff, G . , Ed . ; Simon & Schuster, New York: 1981; entries 20 and 329. 31. M a , J.; Stoter, G . ; Verweij, J. Schellens, J. H . M . Cancer Chemother. Pharmacol. 1996, 38, 391-394. 32. Perezcorona, T.; Madrid, Y . ; Camara, C. Anal. Chim. Acta 1997, 345, 249. 132 Chapter 4 ARSENIC IN THE MEAGER CREEK HOT SPRINGS ENVIRONMENT 4.1. Introduction The Meager Creek hot springs are located north of Pemberton, in British Columbia, Canada. Hot springs are formed when water percolates through permeable rock or fractures, is heated by the earth's crust at depth, and is then driven to the earth's surface by a combination of artesian flow and thermal convection 1. Hot water is able to dissolve minerals over time, so that elevated levels of metals and metalloids are often associated with hot springs. For example, arsenic concentrations in water from hot springs in Yellowstone National Park have been documented to reach 1.6 ppm 2 . Levels of total arsenic reported for a number of Japanese hot springs range from non-detectable to 25 ppm 3. The presence of arsenic in geothermal waters is due to the dissolution of arsenic containing minerals such as arsenopyrite, niccolite, enargite, orpiment, realgar, proustite and other pyrites4, and the most probable form of arsenic in hydrothermal solutions is thought to be arsenite5'6. Pyrite is the most likely host for arsenic at Meager Creek 7. The hot springs at Meager Creek provide an opportunity for studying the environmental chemistry of metal(loid)s such as arsenic. Arsenic is a known poison and carcinogen, and its toxicity is dependent on the chemical form, or species, that it takes. For example arsenobetaine ((CH3) 3As +CH 2COO") can be found in marine animals and mushrooms, and is much less toxic than arsenous acid (As(OH) 3 ) . Some arsenic compounds found in the environment are listed in Table 1.1 (Chapter 1). 133 Very little is known about arsenic speciation in a hot springs environment specifically, although limited knowledge is available about non-marine ecosystems. For example, arsenic speciation was determined in a river in Japan receiving drainage from hot springs8, but samples were not collected from the immediate hot springs environment. Samples analyzed included a green alga, a diatom, freshwater fish, a freshwater prawn, a marsh snail and fly larvae, and all samples contained D M A and T M A when they were analyzed by H G - G C - A A S following alkaline digestion of extracts. These results are inconclusive because the methodology used involves a digestion procedure, which may change the species of arsenic, and an analytical tool that utilizes derivatization and hence does not permit identification of the arsenic species in solution. A series of studies on freshwater and terrestrial plant uptake of radioactive arsenic has been published 9 ' 1 0 ' 1 1 and arsenolipids were found to be formed in aquatic plants, although no conclusive structural elucidation was carried out. In another study five species of halophytes from an estuarine environment were analyzed for arsenic species and content 1 2. The authors claim that compounds such as arsenobetaine, arsenocholine, tetramethylarsonium, T M A O , D M A and an arsenosugar were found in plant extracts and that metabolic synthesis of these compounds is taking place within the plants. The present study was undertaken to determine, semi-quantitatively, arsenic levels and speciation in water and biota at Meager Creek hot springs to extend knowledge about the chemistry of arsenic in freshwater/terrestrial environments7"10. 134 4.2. Experimental 4.2.1. Chemicals and reagents Arsenic standards were obtained as sodium arsenate, Na2HAs04.7H20 (Aldrich), arsenic trioxide, AS2O3 (Alfa), methanearsonic acid, C H 3 A s O ( O H ) 2 (Vineland Chemical), and cacodylic acid, ( C H 3 ) 2 A s O ( O H ) (BDH) , and were dissolved in deionized water to make standard solutions. Extracts of kelp powder (Galloway's, Vancouver, B C ) and Nor i (Porphyra tenerd) of known arsenosugar content 1 3 were used to identify the retention times of arsenosugars; these were verified by comparison to pure arsenosugars generously donated by K . Francesconi and T. Kaise. Arsenobetaine 1 4, arsenocholine1 5, trimethylarsine oxide 1 6 , and tetramethylarsonium iodide 1 7 had been synthesized previously according to standard methods. Methanol ( H P L C grade, Fisher), tetraethylammonium hydroxide ( T E A H , 20% in water, Aldrich), malonic acid (BDH) , concentrated phosphoric acid (Aldrich), ammonium hydroxide ( I M , Fluka), pyridine (Fisher), and formic acid ( B D H ) were used as reagents for mobile phases and extractions. 4.2.2. Sampling Sampling was carried out in November, 1996 and July, 1997. Sample locations are shown in Figure 4.1. Water was sampled by hand into polypropylene bottles that had been acid washed previously. Biota were sampled by hand, stored in Ziploc® bags and kept cool until processing in the lab. There, they were washed thoroughly with tap water to remove soil and other particles, rinsed with deionized (1 Mohm) water, and frozen. Microbial mats were rinsed with only a minimum of deionized water before freezing, to prevent loss of mat composition. The samples were then freeze-dried and pulverized to a fine powder for analysis. 135 Figure 4.1. Map (not to scale) of Meager Creek Hot Springs area showing sampling locations 136 Identification of plants and mushrooms (including bracket fungus) was carried out by using field guide books 1 8 ' 1 9 ' 2 0 . Assistance from M s . O. Lee and Dr . W . B . Schofield (Botany Department, U B C ) , and from M r . James Black (Vancouver Mycological Society) is greatly appreciated in the identification of moss, lichens and mushrooms. 4.2.3. Sample preparation and analysis For the determination of total arsenic content by using I C P - M S , all samples were analyzed in duplicate resulting in <2% standard deviation. Water samples were analyzed directly by I C P - M S ( V G PlasmaQuad, V G Elemental) using R h (10 ppb) as an internal standard. Acid digestions of biota samples were carried out after weighing (0.3 g ± 0.5 mg) the freeze-dried powders into either a 40 m L glass vial or a 500 m L round bottomed flask (RBF) . Concentrated nitric acid (3 mL, doubly distilled in quartz, Seastar, Sidney, B C ) was added to each sample, and additionally, hydrogen peroxide (3 mL, 30% in water, reagent grade, Fisher) was added to each sample in glass vials. The samples in glass vials were heated directly on a hot plate and boiled for 3 hours. The contents of the R B F s were boiled for 2 hours by using a heating mantle and a reflux apparatus2 1 and then cooled. Hydrogen peroxide (3 mL) was added to the R B F s and the solutions were heated for another hour. After all the samples had cooled, the clear solutions remaining were diluted to 25 mL with deionized water and stored until analysis. The acid digests were analyzed by using I C P - M S , with R h (10 ppb) as an internal standard, and by monitoring m/z 75 and 103 for arsenic and rhodium, respectively. I C P - M S parameters are given in Table 4.1. Extractions were carried out by weighing 0.5 to 1 g (+ 0.5 mg) of the freeze-dried powders into 50 m L or 15 mL centrifuge tubes, adding 10-15 mL MeOH/Water (1:1), sonicating for 20 minutes, centrifuging for 20 minutes and decanting the liquid layer into a R B F . Each 137 sample was sonicated and centrifuged a total of 5 times. The decanted extracts for each sample were pooled and rotovapped to near dryness (1-2 mL) and then dissolved in 5 or 10 mL of deionized water. The extracts were filtered through 0.45 um syringe filters (Millipore) and analyzed by H P L C - I C P - M S using the conditions given in Tables 4.1 and 4.2. Data from the I C P - M S were processed by using chromatographic software2 2, and identification of arsenicals in samples was made by comparison of retention times with those of standards by using at least two chromatographic systems. Semi-quantitative concentrations of arsenic compounds were determined by using external calibration curves for each compound corresponding to a matching standard, or to D M A for arsenosugars. Table 4.1. Operation parameters for I C P - M S Specific Conditions Feature Forward radio-frequency power Reflected power Cooling gas flow rate (Ar) Intermediate (auxiliary) gas flow rate (Ar) Nebulizer gas flow rate (Ar) Nebulizer type Analysis mode Quadrupole pressure Expansion pressure 1350 W <10 w 13.8L/min 0.65 L/min 1.002 L/min de Galan Time Resolved Analysis ( T R A ) for H P L C 9 x 10"7 mbar 2.5 mbar 138 Table 4.2. H P L C conditions for arsenic speciation Chromatography Anion exchange Cation exchange Ion-pairing Column Mobile phase Flowrate (mL/min) 1.0 Hamilton PRP-X100 , 150 20 m M ammonium x 4.6 or 250 x 4.6 mm phosphate, p H 6.0 Supelcosil L C - S C X or Whatman S C X Partisil 5, 250 x 4.6 mm G L Sciences O D S , 250 x 4.6 mm 20 m M pyridinium formate, p H 2.7 10 m M T E A H , 4.5 m M malonic acid, 0.1% M e O H , p H 6.8 0.8 4.3. Results and Discussion 4.3.1. Total concentrations of arsenic in water samples The concentrations of arsenic and sampling locations (Figure 4.1) for water samples taken in November 1996 and July 1997 are shown in Table 4.3. These results show no seasonal variations from midsummer to autumn sampling, with the arsenic levels being very similar with respect to both the locations sampled and the sampling date (range of 237-303 ppb). The concentration of arsenic in the hot springs water (average concentration of 280 ppb) is two orders of magnitude higher than in the cold Meager Creek water (5.4 ppb), reflecting the action of hot water on arsenic containing minerals. The cooler temperature observed at location 1 in November was probably a result of cooler air temperatures (-5 to 0 °C) and precipitation in the form of snow. The oxygen concentration ([0 2]) results show that the water was well oxygenated at the source, at outlet 1+2, and at the top of the microbial mats, but more reducing conditions exist under the microbial mats and in the sediments. H P L C - I C P - M S analysis of waters 2 3 showed only the presence of arsenate, except for trace levels of arsenite at location 2. This indicates that i f the arsenic was dissolved initially from minerals as arsenite, oxidation to arsenate took place before expulsion of the water from the source. Arsenate is the species of arsenic expected in most waters 2 4, although others have found up to 80% of total arsenic as arsenite in Hot Creek, California 2 5. 140 Table 4.3. Some physical and chemical characteristics of Meager Creek waters (see Figure 4.1 for sample locations), na = not analyzed. Location Date [As] (ppb) [0 2 ] (ppm) T ( ° C ) p H (SD) a % Source 2 Nov. 1996 237 (8) na na na 1 Nov. 1996 303 (2) b 3.04, 56.7% c 44 6.7 1.08, 20.2% d 4 (geyser) Nov. 1996 288 na na na 8 (Meager Nov. 1996 5.4 na 3 8.3 Creek) Source 2 July 1997 na 2.70, 38.4% 56 6.4 1 July 1997 286 b 0.60, 8.5% d 52 6.3 2 July 1997 277 b 0.75, 10.0% e 41 6.8 1 + 2 outlet July 1997 289 b 4.20, 59.7% 49 6.3 a SD = standard deviation from analysis of duplicate samples; other concentrations are for single samples only. b Also in Feldmann et at23 0 Measurement taken at surface of microbial mat. d Measurement taken below microbial mat. e Measurement taken in sediment below microbial mat. 4.3.2. Total concentrations of arsenic in biota The concentrations of arsenic in acid digested samples of biota sampled at two different times, in November 1996 and in July 1997 are shown in Table 4.4. Sampling locations correspond to numbers on the map of the sampling area in Figure 4.1. 141 Table 4.4. Samples, sampling location (see Figure 4.1), sampling times and arsenic levels (ppm dry weight) (SD for duplicate digestions) in biota samples, B = background Sample Arsenic Sampling Sampling (ppm) time location Top layer, microbial mat 290 (20) N o v 1996 1 Brown microbial mat, 82 (8) N o v 1996 5 Algae 1 (deep green algae) 249 N o v 1996 2 Sedge, Scirpus sp. 7.1 (0.4) N o v 1996 1 Cedar, Thuja plicata 0.96 N o v 1996 3 Moss at geyser, Fumaria hygrometrica 237 (2) N o v 1996 4 Fleabane, Erigeron sp. 14(2) N o v 1996 1,2 Brown lichen, Bryoria sp. 0.30 N o v 1996 6 Yel low lichen, Alectoria sp. 0.12 N o v 1996 6 Orange microbial mat, bottom 59 July 1997 5 Orange microbial mat, top 108 July 1997 5 Algae 2 (green algae) 56 July 1997 5 Sedge, Scirpus sp. 4.5 July 1997 1 Moss at geyser, Fumaria hygrometrica 350 July 1997 4 Moss at stream, Fumaria hygrometrica 91 July 1997 3 Fleabane, Erigeron sp. 3.9 July 1997 1,2 Monkey flower, Mimulus sp. 8.7 July 1997 1,2 Brown lichen, Bryoria sp. 0.30 July 1997 6 Brown lichen, Bryoria sp. (B) 4.8 July 1997 7 Yel low lichen, Alectoria sp. 0.16 July 1997 6 Yel low lichen, Alectoria sp. (B) 0.55 July 1997 7 Cup mushroom, Tarzetta cupularis 0.09 July 1997 7 Fawn mushroom, Pluteus cervinus 0.10 July 1997 7 Bracket fungus, Fomitopsis pinicola <0.07 July 1997 7 142 Microbial mats are found covering the rocks over which the hot water flows from the source into the nearby river. These mats are 1-6 thick and consist of a variety of organisms, usually predominantly bacteria and cyanobacteria but also including fungi and algae (their microbiological makeup is discussed in a later section) 2 6. Their appearance can be described as alternating layers of green, brown and orange with pockets of purple. Strings of deep green material can be found on rocks in areas of faster water flow and these were labeled Algae 1. "Brown microbial mat" refers to brown slime covering the bottom of a cooler stream (about 30° C), and Algae 2 refers to a sample of stringy, absorbent algae collected near location 4. Finally, a thick mat of bright orange was found in July in a stagnant pool of cooler water, and this mat was also composed of layers, with orange on top, brown and green layers, and a white crust on the bottom. A l l algae and microbial mat samples contain high levels of arsenic, ranging from 56-290 ppm (dry weight), and the microbial mat from location 1 contains the highest amount. The lowest amounts of arsenic in these samples are observed in the orange microbial mat (59 and 108 ppm), the brown microbial mat (82 ppm) and Algae 2 (56 ppm), all taken from locations furthest away from the hot springs source, with the exception of Algae 2. Dilution (i.e., snow in November), precipitation (as, for example, Ca 3 (As04)2), absorption by biota, or adsorption of the dissolved arsenic may result in lower levels of arsenic in water farther away from the source, and these lower arsenic levels in the water would result in lower levels in the microbial mat samples. N o seasonal changes were studied for these samples. Moss (Fumaria hygrometrica) was sampled at location 4, a geyser source, the result of drilling by B C Hydro in 1974, and also at location 3, a cooler stream about 200 m away from sources 1 and 2 (See Figure 4.1). A greater arsenic content is observed for the moss from location 4 in July (350 ppm) with respect to that in November (237 ppm), and both these 143 amounts are higher compared with the arsenic content in moss from location 3 (91 ppm). Although the differences have not been statistically validated by taking replicate samples from each location and time, they may be due to the following possibilities. The arsenic content in samples taken from the same location in midsummer and late autumn may reflect seasonal differences in arsenic uptake rates and accumulation. Lower levels of arsenic may have been present in the water at the stream location for the reasons mentioned above, leading to lower levels in the moss taken from the stream. Finally, different levels of submergence of the mosses in the arsenic containing water might cause differences in arsenic concentration. Sedge {Scirpus sp., most likely Olney's bulrush, a species known to grow at Meager Creek Hot springs1) and fleabane (Erigeron sp.) samples contain higher levels of arsenic in the late autumn (7.1 ppm for Scirpus sp. and 14 ppm for Erigeron sp.) compared with the samples taken in summer (4.5 ppm for Scirpus sp. and 3.9 ppm for Erigeron sp.), but again these differences have not been statistically validated. Nevertheless, because the levels of arsenic in the water are the same at both these times, the higher arsenic levels observed in the plant samples from late autumn may indicate that the rate of arsenic accumulation exceeds the rate of arsenic depuration through the growing season. Studies on accumulation of arsenic in plants growing on mine dumps showed a similar trend, where accumulation rates were associated with physiological growth of the plants 2 7. Bracket fungus and mushrooms (see Table 4.4 for Latin names), sampled from the forested area nearby (i.e., not directly adjacent to the hot springs) contain low levels of arsenic (0.10 ppm and lower, Table 4.3). This may indicate that the amount of arsenic being transported via the atmosphere from the hot springs is negligible or very low. Lichens (Bryoria sp. and Alectoria sp.) collected from a tree directly adjacent to a steam vent (location 6) do not contain levels of arsenic that are elevated (0.12-0.30 ppm) with respect to samples from the forest, 144 location 7 (0.55-4.8 ppm), indicating that the lichens are not being impacted by the steam vent. Cedar leaves (Thuja plicata) were sampled directly above a warm stream that drained from the hot springs source into the river. The source of arsenic in the cedar was probably the soil in which the tree was growing, rather than the steam generated by the stream. 4.3.3. Arsenic speciation in biota samples Semi-quantitative amounts of arsenic species in samples from Meager Creek have been estimated and are listed in Table 4.5. The structures, names and abbreviations for arsenic compounds can be found in Table 1.1, Chapter 1. 145 2 a c 3 O C D J 3 i2 8 Q on J 3 <D 1 3 B C/3 C D OO o O .3 x 'o 2 CD £ a < u c <D & O O s on oo c O g C C D 3 o o Q 3 Q O c o •a o S C D ca S i 3 •a u iri <*. s E s 1 s o o H Mjj 7) o V o V o o V 8 <U cN 5 o i—i CO • o o o o « CN (S H H o o o o o o o o o o V V V V V co o (N T-< r - 1 i—1 o o o o o o o o o o o o d © d d o V CO o g o d V 0 0 CN d d o CN CN O O CN i—i i—i rn © © © © d d d d V V V V a> S P O © CN 2 d P a v o oo © d 3 CN d P v o m i-H H O O P o d O V V CN CN o o 0 0 d d CO _J CN rt 55 ^ 0 0 d oo cn co ^ ^ ^ O O CN CN d d d d o v o d o V co CN o o CN CN o o d d V V o © © o d d V V V CN H H O O O o d d V V V ^ CN © n -d d CN u-> I-M' P U CN CN © s - - r CN o r n r- o d d - H - ^ co CO CO O O d d s d ^ <*> s S g ~ O a oo o o V ® © p >T) CO *w CN i—• d © c-d co CD i 4> CO 0\ y CN O d d CN o d v m CO O d V ~ - - § vo vo r - . O O p d d ~T o o co v v CN O S g oo d o vo ON ON > o z ca S S p > o 2 CN VO Ov Ov > 0 0 r-O N O N O N n N O V O O N O N O N S 3 s § S c p •8 0 0 > s z VO o\ O N § 3 S > 2 £ i 1 a\ j J51 O N 3 O N VO O vo ^ ° ^ O N O N RT 2 •5 .a > c S 5 2 a •a 3 i n O N O N O N O N 9 3 O N O N 3 VO 4-T ca S 3 3 'C o i w 0 0 0 0 •a s §5 'S 'S I • 2 oo a ^ 0) CN op cu ca "3 O N O N >> >, 3 3 l —> i—> VO T3 , - O N O § 2 & © 2 §5 0 0 i OH y i u co CN O d "3 o\ O N — ~3 oo oo X •si •S § S Oil o o < 9" c S on h, k, 5 5 r & V 5 « a « a C C C <=> o 5 o CJ ^ ^ ^5 eq ^ o. a. .8 .5 5. 1 "3 O N O N r- l "3 i—> 2 C 5 —> a. a> g S Q . o a, "S3 « p §1 i i o 0 0 s s c3 1 s s a. 3 s e o « 31 ca S 3 S. I 3 u o It 0 0 H 146 4.3.3.1. Algae and microbial mat samples A l l algae and microbial mat samples contain arsenosugars (Table 5), with the orange microbial mat containing the highest amount proportionally (44% of sum of extracted arsenic species). Only arsenosugars X and X I are observed in samples and in most cases the major arsenosugar is arsenosugar X . Sample chromatograms for Algae 1 and extracts used as laboratory standards (from kelp and Nori) are shown in Figure 4.2, and they show the selectivity of the different chromatographic techniques. Figure 4.2a (ion-pairing chromatography) shows the presence of arsenosugar X I and the absence of arsenite. The absence of arsenite allows arsenosugar X to be seen in Figure 4.2c (anion exchange chromatography), and the presence of arsenosugar X I is confirmed in this chromatogram as well. Figure 4.2b (cation exchange chromatography) corroborates the presence of arsenosugar X , well separated from the other arsenic compounds. Another sample chromatogram is illustrated in Figure 4.3, showing the matching retention times for arsenosugar X I when a sample (top layer, microbial mat) is spiked with an extract containing arsenosugar X I (Nori). 147 120 2 4 0 360 4 8 0 6 0 0 720 units) I 4.2b bitrary A s ( V ) / X I A l g a e 1 Nor i 1 -CO-0 ) cz o o X l I i i i i i i i 0 6 0 120 180 2 4 0 300 360 4 2 0 4 8 0 540 6 0 0 0 60 120 180 240 300 3 6 0 T i m e (s) Figure 4.2. Chromatograms for Algae 1 and laboratory standards showing the presence of arsenosugars X and X I . Extracts of Nor i labeled Nor i 1 and Nor i 2 contain predominantly arsenosugar X and X I , respectively. 4.2a. C18 column, 10 m M TEAH/4 .5 m M malonic acid/pH 6.8 mobile phase. 4.2b. Cation exchange column, 20 m M pyridinium formate/pH 2.7 mobile phase. 4.2c. Anion exchange column, 20 m M ammonium phosphate/pH 6.0 mobile phase. 148 As(lll) DMA/As(V) (0 co in tf) 8 Top layer, microbial mat Top layer, microbial mat, spiked with XI 120 240 360 time (s) 480 600 720 Figure 4.3. Chromatograms of a microbial mat extract (top layer, microbial mat) and extract spiked with arsenosugar XI, showing co-elution of arsenosugar XI by using ion-pairing HPLC-ICP-MS (C18 column, 10 mM TEAH/4.5 mM malonic acid/pH 6.8 mobile phase). 149 Arsenosugars have been found in the environment previously in marine algae, which contain, among others, arsenosugars X through XIII , often as the only species of water soluble * 13 28 29 30 * arsenic ' ' ' . In marine phytoplankton Heterosigma akashiwo and Skelotonema costatum (a diatom), arsenosugars X I and XIII were found to be the major compounds in each organism, respectively 3 1. Arsenosugar XIII was the major metabolite produced when another marine diatom, Chaetoceros concavicornis, was exposed to arsenate32. This previous work may indicate that arsenosugars are formed de novo by these lower trophic level marine organisms. The de novo synthesis of arsenosugars by the marine macroalga Fucus spiralis has been postulated to take place, based on the transformation of radiolabeled arsenate in seawater to water soluble organoarsenic compounds within the alga, and subsequent speculation that the compounds were arsenosugars X and X I 3 0 ' 3 3 . Other marine green algae, Dunaliella sp. and Polyphysa peniculus do not form arsenosugars in pure culture; the former reduces arsenate to arsenite3 4 and the latter methylates arsenic species3 5. Arsenosugars appear to be not as common in terrestrial and freshwater environments. The freshwater algae, Chlorella sp. have been studied extensively and the extractable arsenic species after exposure to arsenate is predominantly arsenate, or inorganic arsenic 3 6 ' 3 7 ' 3 8 . Goessler etal.3S, however, found that in addition to arsenate, Chlorella Bohm forms small amounts of an unidentified arsenical, arsenite, M M A and D M A . Comparing the H P L C retention time of the unknown compound to the retention time for arsenosugar X I in our studies, and noting that a similar chromatographic system to ours was used, may allow the unknown compound to be tentatively identified as arsenosugar X I . A commercial sample of the terrestrial soil/freshwater cyanobacterium, Nostoc sp. contains arsenosugar X 1 3 , making up 32% of the total arsenic, but in pure culture Nostoc sp. takes up arsenate as inorganic arsenic, with less than 1% D M A formed 3 9. Likewise, another freshwater cyanobacterium Phormidium sp. contains mostly inorganic arsenic 150 that is not free, but N a O H digestable . A freshwater diatom was found to contain D M A (81%) by using an inconclusive method of speciation8, allowing for the possibility that the D M A found is actually derived from arsenosugars in the diatom. Hence arsenosugars are possibly formed de novo by certain species of marine algae (micro and macro), freshwater cyanobacteria and freshwater algae. Microbial mats have been described as "a heterotrophic and autotrophic community, dominated by cyanobacteria...annealed tightly together by slimy secretions from various microbial components" 4 1. The microbial compositions of mats from Yellowstone National Park and elsewhere have been found to depend on the pH, the temperature, and the H 2 S concentration in the water 2 6. Based on those factors, microbial mats at locations 1 and 2 at Meager Creek possibly contain cyanobacteria species such as Synechococcus lividus, the photosynthetic green nonsulfur bacteria Chlorflexus sp. and the purple bacterium Chromatium tepidum26. However, the mats are almost certainly not homogeneous with respect to time or area and many other bacteria are present, as mentioned earlier. For example, 22 species of bacteria and 1 fungus have been cultured from the microbial mat at location l 4 2 . Therefore, the arsenosugars found in the mats are probably synthesized by cyanobacteria or other bacteria. The high proportion of inorganic species, especially arsenate, in most samples, may imply that most of the arsenate in the water is not metabolized by the microbes making up the mat. Similar behaviour has been observed with bacteria cultured from a marsh area in Yellowknife, N W T 4 j . Arsenite was extracted from the microbial mat sample taken from location 1, which is not surprising considering the reducing conditions imposed by the mat (Table 4.3). This suggests that arsenate-reducing bacteria are present in this mat. The microbial mat from location 1 contains M M A , and the orange microbial mat from location 5 contains D M A (Table 4.5), indicating that methylation is taking place by organisms in these mats. 151 Microbial mats were present in the algae sampling locations, and hence the arsenosugars and other arsenic compounds extracted from algae may be due to uptake from their environment. Therefore although the possibility exists that these algae form arsenosugars, no conclusions can be made about their capability to do so. A difference to note between freshwater and marine arsenosugar formation is the finding of only arsenosugars X and X I in Meager Creek and other terrestrial samples 1 3 ' 3 8, whereas arsenosugars X I I and XIII , as well as arsenosugars X and X I , are common in marine algae. Among marine algae, differences have also been noted; arsenosugars X and X I predominate in Rhodophyta and Chlorophyta, and arsenosugars X I I and XIII are the major compounds in Phaeophyta 3 0. The phylum Phaeophyta is almost exclusively marine and hence the lack of sugars X I I and XIII in the freshwater environment is not surprising, i f these sugars are restricted to Phaeophyta. Microbial mats appear to follow the same chemotaxonomic trend as Rhodophyta and Chlorophyta. However, the reasons for the trends are not clear, since the aglycones (R + groups in the pathway shown in Figure 1.2., Chapter 1) are found as common metabolites in all organisms3 0. 4.3.3.2. Vascular plants (sedge, cedar, fleabane, monkey flower) The identities and amounts of arsenic species associated with an organism probably reflect at least four things: (a) the presence of arsenicals outside the organism, in its food source; (b) the ability of these compounds to enter the organism; (c) the ability, i f any, of the organism to synthesize arsenic compounds and (d) the presence of arsenicals adsorbed onto the outside surface. At this point it is unclear how much these factors influence the speciation of arsenic found in plants. In previous studies, Nissen et al 9 and de Bettencourt et al. 12 have implied that arsenicals other than arsenate found in plants were metabolites and hence synthesized by plants, 152 but de Bettencourt et al. also allowed for the possibility that arsenic speciation in halophytes reflected the arsenic species found in the estuarine waters from which they were sampled 1 2 ' 4 4. In other studies, Catharanthus roseus (periwinkle) was conclusively shown to methylate M M A to D M A in 4 % yield in pure plant tissue culture 4 5. Other authors believe that trace amounts of methylated species in plants are due to uptake from the soil after the compounds are formed by microbial activity in the soi l 4 6 . This theory is supported by studies in which beans and vascular aquatic plants absorbed M M A from soil and water 4 7 ' 4 8 ' 4 9 . The major extractable arsenic species in vascular plants from Meager Creek hot springs are arsenite and arsenate (Table 4.5). A l l the plants, with the exception of cedar, were growing in wet soil that consisted of microbial mats, at the edges of the streams. Although the soil in which the plants were growing was not sampled, it is reasonable to suggest that there may be arsenosugars in the root environments of the plants. Therefore it is interesting to note the lack of arsenosugars in the plants. These results are consistent with some of the studies mentioned above, where inorganic arsenic species are the major water soluble species of arsenic found in plants. Differences are seen between the July and November samples for sedge and fleabane (Figure 4.4). In sedge, higher levels of arsenite and M M A are seen in November, and in fleabane, higher levels of arsenite are seen but slightly lower levels of D M A are seen in November. These observations may suggest that more biological activity is taking place outside the plant, and accumulation of these compounds is taking place, leading to higher levels of arsenite and methyl species inside the plant. However, no seasonal reduction or methylation was evident in the water sampled at any locations, although water analyzed from microbial mats incubated anaerobically over time acquired arsenite and methylarsenic species2 3. Arsenic biomethylation and reduction by plants is a possibility as well, and the metabolism taking place 153 may parallel the increased growth of the plant during the summer. Figure 4.4. Seasonal arsenic speciation in higher plants, sedge (Scirpus sp.) and fleabane (Erigeron sp.), shown as proportions of total arsenic extracted. Interestingly, the major extractable arsenic species in cedar is arsenite (Table 4.5). When pine seedlings, Pinus sp., were allowed to take up radiolabeled arsenate through their roots hydroponically, arsenite was found as the major species in roots and shoots 1 0, which may indicate reduction by the plant. Arsenate was found to be more toxic than arsenite to Catharanthus roseus45. Hence reduction to arsenite by cedar, and by the other vascular plants collected from Meager Creek, may in fact be a detoxification mechanism. The monkey flower Mimulus sp., in addition to containing nearly equal amounts of extractable arsenite and arsenate (2.8-3 ppm dry weight), contains a small amount (0.014 ppm) of tetramethylarsonium ion (Table 4.5, including footnotes to the table). This is a compound that was not found in the microbial mats, in water, or in water after fermentation of microbial mat 2 3. 154 The occurrence of Me4As+ may indicate the presence of a specific microenvironment in which this compound is available to the plant, or an ability of the plant to synthesize Me4As+. Marine sediments cultured aerobically with M M A and D M A released Me4As+ into the culture medium in small amounts 5 0 and tetraalkylated arsenicals (but not Me4As+) were reported to have been found in estuarine waters 4 4 ' 5 1 . Nevertheless, M e 4 A s + has otherwise not been found in pure or mixed culture experiments, in sediments, or in waters; this information together with the lack of the compound in Meager Creek waters may suggest that methylation is indeed taking place in this plant. The formation of Me4As+ is thought to be the final product of the methylation pathway proposed by Challenger, with methylation of trimethylarsine by methyl donors such as S-adenosylmethionine ( S A M ) or methyl halides 2 4. However, normally the reaction sequence stops at or before the formation of M e 3 A s O (or M e 3 A s ) . A n alternative pathway has been suggested, which involves the exchange of the ribosyl group on a precursor trimethylarsonioriboside (Chapter 1, Figure 1.3, intermediate 3.1) for a methyl group 5 2 . The complete absence of the intermediate trimethylarsonioriboside, as well as the absence of its precursor molecule, arsenosugar XIII , in the terrestrial environment, makes this pathway unlikely in terrestrial plants. In summary, the speciation of water soluble arsenic in vascular plants can be generalized as follows: plants may be selective for inorganic arsenic, they may be able to biomethylate arsenic in small amounts and they may detoxify arsenate by reducing it to arsenite, which may be less toxic to them. 4.3.3.3. Moss Arsenosugar X was detected in all moss samples (Table 4.5). Microbial colonies were observed in and around the moss samples, and despite the washing procedure, most likely contaminated the samples. The sampling methods used in the present study are not capable of 155 differentiating between the contribution from microbial mat components interspersed throughout the moss sample and the contribution from arsenosugar uptake by the moss from its environment, or of metabolism within the moss. The normalized concentrations of arsenic species in moss are shown in Figure 4.5. Seasonal variations (from July to November) include a greater proportion of arsenosugar X in November, and the presence of methyl species in July. The most likely reason for these differences is that the microbial population in the moss environment changes throughout the growing season. The same reasoning can be used to explain the differences in the moss samples taken from different locations. For example, the moss sampled at the geyser location (location 4) appears to contain less arsenosugar, proportionally, than the moss sampled at the cooler stream location (location 3). This larger proportion of arsenosugar X as well as the presence of arsenite is also seen in the orange microbial mat taken from a cooler location (Table 4.5). Therefore the microbial community in the vicinity of the moss sample at the cooler stream location may resemble a microbial mat at a cooler location, which may consist of more arsenic-metabolizing organisms. 156 Nov moss (L4) July moss (L4) July moss (L3) Figure 4.5. Seasonal and spatial arsenic speciation in moss (Fumaria hygrometrica) shown as proportions of total arsenic extracted, L = location. July and November moss are from the geyser location (L4) and the second July moss is from the stream location (L3) . 4.3.3.4. Fungi including lichens The levels of arsenic compounds in all fungi samples (Table 4.5) are at or close to the limit of detection for the H P L C - I C P - M S methods used. Brown and yellow lichens both contain arsenosugar X L as does the yellow lichen sampled from a location not expected to be impacted by the hot springs (location 7). Pixie cups contain trace levels of arsenosugar X . The presence o f arsenosugars in these samples is very likely a result of synthesis by the indigenous algae or cyanobacteria living symbiotically with the fungus, making up the lichen, since the organisms were relatively isolated, and not exposed to the microbial mats with their sugar-synthesizing organisms. The organisms associated with both pixie cups (Cladonia sp.) and yellow lichen (Alectoria sp.) are green algae of the genus Trebouxia53. O f course, the possibility cannot be discounted of sugars being present due to uptake or adsorption from other organisms living in the immediate surroundings of the lichens, or as a result of metabolism by the fungus. The results for the two mushrooms sampled add to the current knowledge of arsenic speciation in mushrooms. Inorganic species are the major species found, as well as D M A in minor amounts (Table 4.5). Arsenosugar X was detected in Tarzetta cupularis, but in trace amounts; verification of this result, especially in specimens containing higher levels of arsenic, would be very interesting. A small amount of an arsenosugar was reported in another mushroom, Laccaria amethystina, but the authors considered additional chromatographic confirmation to be necessary5 4. Very low levels of water soluble arsenic are present in these fungi. 4.3.3.5. Extraction efficiency for arsenic species The sums of arsenic species extracted are low for all samples analyzed. The % amounts of extracted arsenic with respect to the total, obtained by dividing the sum o f species in Table 4.5 by the total arsenic in Table 4.4, are shown in Table 4.6. This calculation does not take into account the amount of arsenic that may have been extracted but was not observable by using these chromatographic methods. In past studies, this amount has been observed to be significant in some samples, even when using the method of standard additions for quantification5 5. Analytical problems have been reported when significant levels of sodium and potassium were present in samples; that is, a suppressed chromatographic signal for arsenobetaine resulted from the co-elution of sodium and potassium ions during cation exchange H P L C - I C P - M S 5 6 . The suppression of the signal for earlier eluting arsenic species (e.g., anionic species and arsenosugar X ) is possible as well, although such an effect was not reported 5 6. 158 Table 4.6. Percent amounts of arsenic extracted. Sample % Arsenic extracted Top layer, microbial mat, N o v 1996 31 Brown microbial mat, N o v 1996 2.4 Algae 1 (deep green algae), N o v 1996 1.9 Sedge, Scirpus sp., N o v 1996 13 Cedar, Thuja plicata, N o v 1996 21 Moss at geyser, Fumaria hygrometrica, N o v 1996 0.6 Fleabane, Erigeron sp., N o v 1996 47 Brown lichen, Bryoria sp., N o v 1996 28 Yel low lichen, Alectoria sp., N o v 1996 41 Orange microbial mat, bottom, July 1997 5.4 Orange microbial mat, top, July 1997 1.7 Algae 2 (green algae), July 1997 6.7 Sedge, Scirpus sp., July 1997 31 Moss at geyser, Fumaria hygrometrica, July 1997 1.1 Moss at stream, Fumaria hygrometrica, July 1997 1.9 Fleabane, Erigeron sp., July 1997 47 Monkey flower, Mimulus sp., July 1997 67 Brown lichen, Bryoria sp., July 1997 10 Yellow lichen, Alectoria sp., July 1997 19 Yellow lichen, Alectoria sp., July 1997 (B) 19 Cup mushroom, Tarzetta cupularis, July 1997 70 Fawn mushroom, Pluteus cervinus, July 1997 63 Extractable amounts of arsenic range from 0.6 % for moss to 70 % for the mushroom Tarzetta cupularis. From vascular plants, 13-67% of total arsenic was extracted (mean of 38%), from moss, 0.6-1.9% of total arsenic was extracted (mean of 1.2 %), and from fungi, 10-70% of total arsenic was extracted (mean of 36%). The extraction method used has been successful for 159 marine plants and animals ' and is assumed to extract water-soluble species, yet it appears to be insufficient for the extraction of these samples. The levels of inorganic arsenic may be underestimated, because in other studies the highest amounts of inorganic arsenic species from terrestrial samples were extracted with water alone 5 9. Insufficient extraction may also result from the presence of arsenic compounds that are nonpolar and not soluble in methanol/water (1:1). In other studies where the presence of arsenolipids was postulated, an extraction technique with hot ethanol was used 9 ' 1 0 ' 1 1 . Other researchers have used a sequential extraction technique consisting of Soxhlet extraction with 80% methanol, cold 5% chloroacetic acid, warm 75% ethanol and hot 5% chloroacetic acid to extract all but 20% of arsenic from wheat seeds60. These extraction techniques are likely to extract compounds that are more nonpolar in nature. Additionally, prior freeze-drying of samples has been recently found to result in lower extraction efficiencies, although the reasons for this are unclear 5 5. Arsenic that is not extracted by methanol/water (1:1) might be tightly bound to lipids; to cell wall components of plants, including insoluble cellulose, calcium or magnesium pectates, or lignin; and to chitin or other cell components of fungal samples. L o w extraction efficiencies are associated with microbial mats and algae (1.7-31%, mean of 18%), a result also seen with Nostoc sp. (34%) 1 3 and possibly with Phormidium, where the arsenic in the cyanobacterium was thought to be not free and possibly bound up in a non-water soluble form 4 0 . Metals (possibly including arsenic) can bind to bioflocculants secreted by microbial mats 4 1, and this may limit extraction efficiencies for arsenic. More specifically, arsenate may coprecipitate with insoluble minerals as a result of microbial mat biomineralization. Evidence exists for the formation of hydrated iron and manganese oxides, and iron and aluminum silicates on bacterial cells in freshwater microbial mats 6 1 ' 6 2, which are minerals known to co-precipitate with arsenate. There is also evidence of ferric arsenate precipitation as a result of 160 bacterial action 4.3.4. Summary Arsenosugars are apparently formed by cyanobacteria/bacteria, possibly by algae in the lichens Cladonia sp., Bryoria sp. and Alectoria sp., and possibly by the fungus Tarzetta cupularis. 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Chem. 1990, 4, 181-190. 29. Shibata, Y . ; Jin, K . ; Morita, M . Appl. Organomet. Chem.1990, 4, 255-260. 30. Francesconi, K . A . ; Edmonds, J. S. Adv. Inorg. Chem. 1997, 44, 147-189. 31. Shibata, Y . ; Sekiguchi, M . ; Otsuki, A . ; Morita, M . Appl. Organomet. Chem. 1996, 10, 713-719. 32. Edmonds, J. S.; Shibata, Y . ; Francesconi, K . A . ; Rippingale, R. J.; Morita, M . Appl. Organomet. Chem. 1997,11, 281-287. 33. Klumpp, D . W. ; Peterson, P. J. Mar. Biol. 1981, 62, 297. 34. Takimura, O.; Fuse, H . ; Murakami, K . ; Kamimura, K . ; Yamaoka, Y . Appl. Organomet. Chem. 1996, 10, 753-756. 35. Cullen, W . R ; Harrison, L . G . ; L i , H . ; Hewitt, G . Appl. Organomet. Chem. 1994, 8, 313. 163 36. Maeda, S.; Ohki, A . ; Kusadome, K . ; Kuroiwa, T.; Yoshifuku, I.; Naka, K . Appl. Organomet. Chem. 1992, 6, 213-219. 37. Kuroiwa, T.; Ohki, A ; Naka, K . ; Maeda, S. Appl. Organomet. Chem. 1994, 8, 325-333. 38. Goessler, W. ; Lintschinger, J.; Szakova, J.; Mader, P.; Kopecky, J.; Doucha, J.; Irgolic, K . J. Appl. Organomet. Chem. 1997, 11, 57. 39. Maeda, S.; Mawatari, K . ; Ohki, A ; Naka, K . Appl. Organomet. Chem. 1993, 7, 467-476. 40. Maeda, S.; Fujita, S.; Ohki, A . ; Yoshifuku, I ; Higashi, S.; Takeshita, T. Appl. Organomet. Chem. 1988, 2, 353-357. 41. Bender, J.; Lee, R. F.; Philips, P. J. Industrial Microbiology 1995, 14, 113-118. 42. Polishchuk, E . ; Liao, T.; Koch, I., University of British Columbia, unpublished results. 43. Koch, I.; Liao, T.; Polishchuk, E . , University of British Columbia, unpublished results. 44. de Bettencourt, A . M . ; Florencio, M . H . ; Duarte, M . F. N . ; Gomes, M . L . R.; Vilas-Boas, L . F. C. Appl. Organomet. Chem. 1994, 8, 43-56. 45. Cullen, W. R.; Hettipathirana, D . ; Reglinski, J. Appl. Organomet. Chem. 1989, 3, 515-521. 46. Pyles, R. A ; Woolson, E . A . J. Agric. Food Chem. 1982, 30, 866-870. 47. Sachs, R. M . ; Michael, J. L . 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Goessler, W. ; Kuehnelt, D . ; Schlagenhaufen, C ; Slejkovec, Z . Irgolic, K . J. J. Anal. At. Spectrom. 1998, 13, 183-187. 57. Shibata, Y . ; Morita, M . Appl. Organomet. Chem. 1992, 6, 343-349. 58. Alberti, J.; Rubio, R.; Rauret, G. FreseniusJ. Anal. Chem. 1995, 351, 420-425. 59. Kuehnelt, D . ; Goessler, W. ; Schlagenhaufen, C ; Irgolic, K . J. Appl. Organomet. Chem. 1997, 11, 859-867. 60. Domir, S. C ; Woolson, E . A . ; Kearney, P. C ; Isensee, A . R. J. Agric. Food Chem. 1976, 24, 1214-1217. 61. Tazaki, K . Clays Clay Min. 1997, 45, 203-212. 62. Konhauser, K . O.; Fyfe, W . S.; Ferris, F. G . ; Beveridge, T. J. Geology 1993, December, 1103-1106. 63. LeBlanc, M . ; Achard, B . ; Othman, D . B . ; Luck, J. M . ; Bertrand-Sarfati, J.; Personne, J.Ch. Appl. Geochem. 1996, 11, 541-554. 64. Robinson, B . ; Outred, H . ; Brooks, R.; Kirkman, J. Chem. Speciation Bioavail. 1995, 7, 89-96. 165 Chapter 5 ARSENIC IN THE YELLOWKNIFE ENVIRONMENT 5.1. Introduction Yellowknife is located on Great Slave Lake, in the Northwest Territories, Canada. A major industry in the city is gold mining, and two gold mines, the Royal Oak Giant Mine and the Miramar Con Mine, are presently in operation. The gold in the mined ore is associated with arsenopyrite (FeAsS), and hence arsenic waste is generated during the smelting operation. For example, when milling began at Giant Mine in 1948, aerial emissions of 7.3 tonnes as arsenic trioxide per day were released1. Currently, aerial emissions of only 1.7 tonnes as arsenic trioxide per year are released from Giant Mine, and 0.5 tonnes of arsenic per year are discharged with the effluent into Baker Creek. The proximity of the two gold mines to the city of Yellowknife has allowed researchers to study the effects of mining and arsenic on the surrounding environment. Arsenic chemistry has been studied to some extent in Pud, Kam, Meg, Keg and Peg watershed areas, which receive the drainage from Con Mine waste. Arsenic mobility was found to be controlled by remobilization of historically contaminated sediments rather than by present day mining practices2. Waters, sediments and porewaters were analyzed for inorganic and methyl arsenic species3 and methylarsenic (III) compounds were discovered although they were not identified. Macrophytes (aquatic plants) were analyzed for total arsenic4 and concentrations were found to be elevated compared with specimens collected from uncontaminated areas. Toxic effects of reduced plant health and lack of biodiversity were observed in areas of high arsenic content. 166 Anaerobic microbes isolated from a sediment core from Kam Lake were found to methylate arsenic to monomethylarsonic acid ( M M A ) , dimethylarsinic acid ( D M A ) , trimethylarsine oxide ( T M A O ) , methylarsenic (III) compounds, arsine and trimethylarsine5. As discussed in the previous chapter, little is known about arsenic speciation in biota in the freshwater and terrestrial environments. The high levels of arsenic in the Yellowknife area allows us to study the forms that arsenic takes in the available biota. A s well, these results can be compared with those obtained from Meager Creek. 5.2. Experimental 5.2.1 Chemicals and reagents Arsenic standards were obtained as sodium arsenate, N a 2 H A s 0 4 . 7 H 2 0 (Aldrich), arsenic trioxide, A s 2 0 3 (Alfa), methanearsonic acid, C H 3 A s 0 ( 0 H ) 2 (Vineland Chemical), and cacodylic acid, ( C H 3 ) 2 A s 0 ( 0 H ) ( B D H ) were dissolved in deionized water to make standard solutions. Extracts of kelp powder (Galloway's, Vancouver, B C ) and Nor i (Porphyra tenera) of known arsenosugar content6 were used to identify the retention times of arsenosugars; the retention times were then verified by comparison to those obtained from pure arsenosugars generously donated by K . Francesconi and T. Kaise. Arsenobetaine7, arsenocholine8, trimethylarsine oxide 9, and tetramethylarsonium iodide 1 0 had been synthesized previously according to standard methods. Methanol ( H P L C grade, Fisher), tetraethylammonium hydroxide ( T E A H , 20% in water, Aldrich), malonic acid (BDH) , concentrated phosphoric acid (Aldrich), ammonium hydroxide ( I M , Fluka), pyridine (Fisher), and formic acid ( B D H ) were used as reagents for mobile phases and extractions. 5.2.2. Sampling Sampling was carried out in June and August of 1997. Sample locations are shown in Figure 5.1a and 5.1b. Water was sampled by hand into polypropylene bottles that had been previously acid washed. Some samples were split and one of the aliquots was filtered through 1 um cellulose nitrate filters (Sartorius) before freezing. Sediment samples were collected by using a trowel and they were frozen in Ziploc® bags. They were air dried over a period of two days, then ground in a mortar and pestle. Particles that passed through a 60 mesh sieve were subsequently extracted. Most biota were sampled by hand and all were stored in Ziploc® bags and kept cool until processing in the lab. There, they were washed thoroughly with tap water to remove soil and other particles, rinsed with deionized (1 Mohm) water, and frozen. Mussels were sampled by hand as well as by using an Eckman grab, and a different freshwater species was collected from Campbell River for comparison. They were shelled before freezing, as were large specimens of snails. Smaller specimens of snails(< 0.5 cm in length) were frozen and processed whole. Fish were caught by using a gillnet, and they were gutted before freezing. Roe was removed from one fish and this was kept and frozen for analysis. The fish were thawed at a later date and filleted and skinned to obtain muscle samples, which were refrozen. Pond scum, or algal/microbial mats of unknown composition, sampled from a puddle beside Baker Creek (location 4) were washed with a minimum of deionized water to minimize sample loss before freezing. A l l frozen biota samples were then freeze-dried and pulverized to a fine powder for analysis. 168 Figure 5.1a. Map of Royal Oak Giant mine property and surrounding area, with sample locations. Figure 5.1b. Map of Yellowknife, showing Royal Oak Giant Mine and Miramar Con Mine, and sampling locations. 170 Mussels and snails 1 1, plants 1 2 and mushrooms 1 3 ' 1 4 were identified by using field guide books. Assistance from O. Lee, Dr. W. B . Schofield and Julie Oliveira (Botany Department, U B C ) is greatly appreciated in the identification of moss, lichens and algae. We are very grateful also to Chris Ollson (Royal Military College) for fish identification, M i k e Fournier (Yellowknife, N W T ) for assistance with plant identification, and James Black (Vancouver Mycological Society) for help with mushroom identification. 5.2.3. Sample preparation and analysis For the determination of total arsenic content by using I C P - M S , all samples except water samples were analyzed in duplicate resulting in <2% standard deviation. Water samples were analyzed only once directly by I C P - M S ( V G PlasmaQuad, V G Elemental) using R h (10 ppb) as an internal standard. Ac id digestions of biota samples were carried out after weighing (0.3 g ± 0.5 mg) the freeze-dried powders into either a 500 mL round bottomed flask (RBF). Concentrated nitric acid in a volume of 3 mL (doubly distilled in quartz, Seastar, Sidney, B C ) was added to each sample. The samples were boiled for 2 hours by using a heating mantle and a reflux apparatus1 5 and then cooled. Hydrogen peroxide (3 mL, 30% in water, reagent grade, Fisher) was added to the R B F s and the solutions were heated for another hour. After all the samples had cooled, the clear solutions remaining were diluted to 25 mL with deionized water and stored until analysis. The acid digests were analyzed by using I C P - M S , with R h (10 ppb) as an internal standard, and by monitoring m/z 75 and 103 for arsenic and rhodium, respectively. I C P - M S parameters are given in Table 5.1. Extractions were carried out by weighing 0.5 to 1 g (± 0.5 mg) of the freeze-dried powders into 50 mL or 15 mL centrifuge tubes, adding 10-15 mL MeOH/Water (1:1), sonicating for 20 minutes, centrifuging for 20 minutes and decanting the liquid layer into a R B F . Each 171 sample was sonicated and centrifuged a total of 5 times. The decanted extracts for each sample were pooled and rotovapped to near dryness (1-2 mL) and then diluted to 5 or 10 mL with deionized water. Fish, fish roe, mussels, snails, and oyster tissue certified reference material ( C R M ) 1566, obtained from NIST , were digested with protease, based on methods for enzymatic digestion 1 6 ' 1 7 ' 1 8 . A n accurately weighed 1 g (±0.5 mg) sample was combined with 0.02-0.05 g of protease (Type VIII, N o . P-5380, Sigma) in a plastic 50 mL centrifuge tube. Ammonium carbonate ( B D H ) buffer at a concentration of 0.1 M and p H 7.2 (adjusted with nitric acid) was added and the tube was sealed and vortexed. The samples were shaken for 4 hours at 37 °C, then centrifuged, and the supernatant was diluted to 25 mL with deionized water. To determine the extent of arsenic solubilization by the protease, oyster tissue samples were extracted with buffer alone (no protease) and using the same procedure as that described for protease digestions. Water samples, extracts and protease digestions (PDs) were filtered through 0.45 um syringe filters (Millipore) and analyzed by H P L C - I C P - M S using the conditions given in Tables 5.1 and 5.2. Data from the I C P - M S were processed by using chromatographic software1 9, and identification of arsenicals in samples was made by comparison of retention times with those of standards by using at least two chromatographic systems. Semi-quantitative concentrations of arsenic compounds were determined by using external calibration curves for each compound corresponding to a matching standard, or to D M A for arsenosugars. 172 Table 5.1. Operation parameters for I C P - M S Specific Conditions Feature Forward radio-frequency power Reflected power Cooling gas flow rate (Ar) Intermediate (auxiliary) gas flow rate (Ar) Nebulizer gas flow rate (Ar) Nebulizer type Analysis mode Quadrupole pressure Expansion pressure 1350 W <10W 13.8 L/min 0.65 L/min 1.002 L/min de Galan Time Resolved Analysis ( T R A ) for FfPLC 9 x 10"7 mbar 2.5 mbar Table 5.2. H P L C conditions for arsenic speciation Chromatography Column Mobile phase Flowrate (mL/min) Anion exchange Cation exchange Ion-pairing Hamilton PRP-X100 , 150 20 m M ammonium x 4.6 or 250 x 4.6 mm phosphate, p H 6.0 Supelcosil L C - S C X or Whatman S C X Partisil 5, 250 x 4.6 mm G L Sciences O D S , 250 x 4.6 mm 20 m M pyridinium formate, p H 2.7 l O m M T E A H , 4.5 m M malonic acid, 0.1% M e O H , pH6 .8 1.0 or 1.5 1.0 0.8 5.3. Results and Discussion Arsenic compounds were identified by comparing retention times of arsenic compounds in samples with those of standard compounds. I f the retention time for an arsenic compound in a sample was the same as that for a standard compound, the arsenic compound was concluded to have the same identity as the standard compound. If the presence of arsenobetaine or cationic species (such as T M A O , arsenocholine, and Me4As+) was indicated after analysis with anion exchange H P L C - I C P - M S , the identities of such species were confirmed by analysis with cation exchange H P L C - I C P - M S method. Cation exchange chromatography was also used to confirm the presence of arsenosugar X . This analytical method separates the aforementioned peaks from other peaks that co-elute with them on other chromatographic systems. The identities of arsenosugars were confirmed by using ion pairing chromatography. Some samples were spiked with standards to confirm that retention times of arsenic compounds did not depend on matrix. The primary purpose of the present study was to determine the identity of detectable arsenic species that could be extracted by using MeOH/water (1:1) extraction. The amounts of the arsenic species found were calculated semi-quantitatively, to determine approximate levels of the compounds (i.e., major or minor components). Standard deviations shown in Table 5.6 and for subsequent results were calculated with quantities obtained from analysis on the different chromatographic systems, where applicable. In some cases, only one number was obtained i f the better sensitivity and selectivity of a chromatographic system allowed the species to be seen on one system but not another. The relative standard deviation, on average, is estimated to be about 30%. 5.3.1. Water and sediment samples Water and soil were sampled in a few locations from which biota were sampled to ascertain the arsenic speciation and concentrations in the surrounding aqueous environment of these biota. Total arsenic concentrations and speciation results for waters, and estimated water soluble species (extracted with MeOH/water 1:1) for soils are summarized in Table 5.3. Table 5.3. Concentrations of total arsenic and arsenic species in water samples (ppb) and soil extracts (ppm) (SD) a . Total concentrations were determined by I C P - M S as described in section 5.2.3. "Trace" amount is 1 ppb (i.e., at detection limit); nd = not determined. Sample (location) As (III) A s ( V ) M M A D M A Sum of species Total _ J 2 C P J ^ M S ) _ Pore water (1) 296 6.2 <1 29 331 980 Surface water (1) 2.5 98 <1 <1 101 140 (30)+ Surface water (2) <1 126 <1 <1 126 nd Surface water (3) <1 184 <1 <1 184 195 (3)J Standing water (4) 1.4 350 (40)+ <1 trace 352 480 Con effluent (14) <1 5.3 <1 <1 5.3 32 (26)t Surface water (10) <1 20.3 2.5 <1 23 68 Surface water (11) trace 26 (5)J 2.5 (0.8)J <1 30 54 (4)t Sediment extract (1) 19.1 0.65 <0.05 <0.05 19.8 nd Sediment extract (2) 14.4 2.0 <0.05 <0.05 16.4 nd Sediment extract (3) 10.8 2.8 <0.05 <0.05 13.6 nd a SD = Standard deviation, based on duplicate analysis at two different dilutions* or analysis of duplicate samples J. In all surface waters the major arsenic compound is arsenate. Minor amounts of methyl species are observed in water samples from Niven Lake (locations 10 and 11) and from location 175 4, which were areas of prolific plant and algal growth. Only small amounts of arsenite are present in some samples, except for the pore water squeezed from a sediment in the Baker Creek marsh, which contains arsenite as the major compound. This indicates strongly reducing conditions in the sediment. Not all o f the arsenic was accounted for in some samples by using H P L C - I C P - M S analysis; the largest discrepancies are observed for the pore water sample, Con effluent, and Niven Lake waters (locations 10 and 11). Species such as methylated arsenic(III)thiols, o f the form ( C H 3 ) n A s m ( S R ) 3 - n (n=l,2,3), as well as those involving binding of arsenic to colloidal organic matter such as humic and fulvic substances, were postulated to be present in waters from Yellowknife in a previous study3. The H P L C behaviour for such arsenic species is unknown and they might not be detectable by using the H P L C - I C P - M S methods in this study. The concentrations of arsenic species in Table 5.3 are in ppm dry weight of sediment. The major extractable species from sediments is arsenite, which probably reflects the insolubility of arsenate in the sediments, and it may reflect reducing conditions as well. I f the MeOH/water technique can be used to approximate the species that are water soluble, then it appears that arsenite is the major species available to organisms growing in the sediment. 5.3.2. Freshwater fish Although arsenic speciation has been studied in some detail for marine fish, very little information is available about the forms of arsenic in freshwater fish. Because of this lack of information, a study was conducted to examine arsenic in fish from Yellowknife. It was also of interest to see i f higher levels of arsenic (compared with published background levels) would be present in the fish as a result of their exposure to higher levels of arsenic in the water and sediments. The sampling area chosen was the Baker Creek outlet in Yellowknife Bay (location 176 9, Figure 5.1a). N o fish are observed in Baker Creek itself after the mine discharge period begins each summer, although the lack of fish in Baker Creek is thought to be due to elevated levels of ammonia rather than metals 2 0. Four different fish species were caught: whitefish (Coregonus clupeaformis), sucker (Catostomus commersoni), walleye (Stizostedion vitreum) and pike (Esox lucius). Fish muscle was chosen as the part of the fish to be analyzed because of its significance to consumers. 5.3.2.1. Total arsenic in fish Some physical characteristics, the moisture content, and the arsenic content in each fish sampled are shown in Table 5.4. For most of these and other samples, acid digestions were carried out once, and analyzed in duplicate by using I C P - M S . Relative standard deviations obtained from duplicate analyses were less than 2% in all cases. When sucker 1, pike 2 and whitefish 3 were digested in duplicate, the average relative standard deviation between digestions was less than 5%. Standard deviations for other samples may be estimated to be 5% of the total arsenic concentration. Total arsenic concentrations have previously been found to be much higher in marine fish than in freshwater fish, and the present results confirm this observation. The highest concentration of arsenic in this study is 3.1 ppm dry weight in a whitefish, whereas arsenic concentrations in marine fish range from 3.5 ppm dry weight for mackerel to 196 ppm dry weight for plaice collected from uncontaminated areas16. Converting these arsenic concentrations to fresh weight concentrations (using [As]/R where R is fresh weight/dry weight in Table 5.4) gives concentrations ranging from 0.07 ppm for whitefish 3 to 0.72 for whitefish 1. 177 Table 5.4. Total arsenic concentrations in fish from location 9. Fish Sex Weight (g) Moisture content in % (R) a Arsenic concentrati< (ppm dry w d g j i t ) j ^ l Whitefish 1 Male 846 " " " " 7 7 ( 4 3 3 ) 3.1 Whitefish 2 Female 586 76 (4.24) 0.84 Whitefish 3 Female 1453 74 (3.88) 0.28 (0.02) Whitefish 3 roe — 137 0.25 Sucker 1 Male 656 78 (4.65) 1.24 (0.01) Sucker 2 Male 457 81 (5.35) 0.98 Walleye 1 Male 386 79 (4.75) 0.46 Walleye 2 Male 507 77 (4.37) 0.85 Pike 1 Male 628 79 (4.72) 1.30 Pike 2 Male 988 80 (5.08) 1.40 (0.09) a Moisture content was calculated by using Q3.-l)/R*100%; R = fresh weight/ dry weight. b SD = Standard deviation, based on duplicate acid digestions. These numbers can be compared with reported arsenic concentrations in freshwater fish sampled from pristine locations, ranging from 0.025 ppm fresh weight for pike to 0.132 ppm fresh weight for white sucker 2 1 ' 2 2 . The concentrations of the fish sampled in this study are lower than those found in rainbow trout and freshwater smelt purchased from a Japanese market (1.46 and 1.08 ppm fresh weight) 2 3. Most likely the levels of arsenic in Yellowknife fish are not elevated compared with fish analyzed in previous studies 2 1 ' 2 2 ' 2 3, but more samples should be analyzed to confirm this. The fish analyzed in this study all contain levels below a maximum permissible concentration of arsenic in fish for human consumption, set by the Australian National Health and Medical Research Council, o f 1.14 mg/kg (ppm) fresh weight 2 1. The sample size from this study is too small to draw any correlations between physical characteristics and arsenic concentration. However, for sucker, walleye and pike, the arsenic 178 concentration appears to increase with increasing fish size, a trend that has been documented previously 2 4. This does not appear to be the case for whitefish. This trend may not be observed for every species offish, and there may be differences based on sex. The concentration of arsenic in the roe is approximately the same as the fish from which it was taken. The roe was very fatty and difficult to digest, and care had to be taken to keep the rate of heating slow to prevent charring. In yellowtail flounder Pleuronectes ferruginea, the arsenic concentrations in spawned eggs were significantly lower than those found in developing gonads and fish muscle 2 5, an observation that contrasts with the results found in this study (i.e., that the concentrations are the same). Clearly, more studies are necessary to determine i f any trends are observable for arsenic concentrations in gonads and muscle for freshwater fish. 5.3.2.2. Arsenic speciation in fish Enzymatic digestion has been used previously for fish and shellfish samples. Branch et al}6 used trypsin, a protease, to disrupt the lipid-protein membrane in fish samples and release the cell contents. They found that trypsin digestion gave greater recoveries for arsenic from a certified reference material, D O R M - 1 (dogfish muscle) and whiting; lower recoveries for plaice, mackerel and a lemon sole specimen; and similar results for cod, when compared with methanol/chloroform (2:1) extraction. Forsyth et a / . 1 7 ' 1 8 used crude protease and lipase to digest samples for tin and lead speciation and observed reduced matrix effects during analysis, effective release of organotin analytes from marine food matrices, and adequate recovery of alkyllead standards. Oyster tissue and dogfish muscle C R M s were digested with a combination of protease and lipase to yield quantitative recovery of selenium 2 6. It was predicted in this work that i f recoveries were similar for enzymatic digestion, using a non-specific protease, compared with acid digestion to determine total arsenic in a C R M , then 179 protease digestion might be a suitable method for releasing arsenic from a protein-rich sample matrix, as well as other elements such as antimony. The results obtained when total arsenic was determined in oyster tissue C R M (NIST 1566) in protease digestions, acid digestions, and aqueous (buffer) extracts are shown in Table 5.5. About 70% of arsenic is recovered from aqueous (buffer) extraction alone, whereas 100% of arsenic is released from the matrix using protease digestion. Higher levels of antimony are present after using the protease digestion, although all values obtained are significantly higher than the non-certified value (0.01 ppm). The reasons for this are unclear, and more studies should be carried out to obtain a more reliable value for the acid digestion procedure to use for comparison. Protease digestion was subsequently used for fish, shellfish and snail samples in an attempt to maximize extraction of other elements, as well as arsenic. Table 5.5. Comparison of arsenic concentrations (SD) a by using protease and acid digestion methods for oyster tissue (NIST 1566). The certified concentration of arsenic is 14.0 ± 1 . 2 ppm, and the non-certified value for Sb is 0.01 ppm. Digestion procedure Concentration of As (ppm) Concentration of Sb (ppm) Acid digestion 14.0 (0.2) Protease digestion, p H 7.2 13.8 (0.2) Buffer extraction, p H 7.2 11.4 (0.3) 0.3 (0.2) b 0.22 (0.08) 0.17 (0.07) a SD = standard deviation, obtained from 3 replicate extractions. b SD for this sample was obtained from 2 replicate digestions. 180 i i (3 ^ 3 Q i © V ° ? 1 eej C > BO ON. c o o o .a 2 a 3 O Q 3 X « .2? 73 CO q c/5 o o. o a x I 9 § 4J C o s o U 3 n H X) Q S c o •a o B <a T3 H - H O i w O a> ' 3 . I oo In5 "8 2 E ••o I X X >- >- >-> >- X X >< >-o o\ ON ON vo s CO r- o HH d *-H <N o VO ts o jo o V o o o o d o o d d V m X O o O O ugar V-H r—1 ugar o o o in CO ugar d o d *-H 0 0 V V V O d 0 0 ; co • © co ,052 (0.00 vo o in CN o vo" o 0 0 o o co • — -r-o lo O ,052 (0.00 d o d d d d © o d -—- d •—• : 0 0 oo CN ,052 (0.00 in CN co CN VO CN o oo CN CN VO CN I d o o o o o d o © d © © © CN CN CN ON in *-H ON VO © CO d CN in o d vo o r-o VO CN d oo UO m d m r-© o CN O d o CN CN CO i-H CN d •n 0 0 in >n d 3 0 0 d vo 3 d s d O hi o O O O. o O ci a d ci ci ci V V V V V V ^ s ^ ^ CO VO o VO o d O d o d d d CO ON •o o. CN OV CN ON I d o d O o o 1 v d V d o" d o 0) CJ C S i s CN O 0 0 d CO CO d d 3 d 0 0 co g O O co sa d d P hi v v o CN O d o d v o o V o d v la a a jCU cu a. CN CO 1 J3 J3 : &o V5 V5 IS IS 1 B Oi h i 12 1^ m O a a. o d v ca hi o d v o d v a CH O d v CO O O d v o d v CO O O d o o o o d d V V V o d o d o d v S a a a ^ CU CU CH CN CN VH w H* CL) <u a) CD -4 -4 _H CJ CJ o O 3 3 3 3 0 0 on 0 0 0 0 co CU >. cu 3 CU 3 CU o o d d V V a CH CN CN cu cu cu cu 2 3 ° - ^ H cu c/3 ca 1 8 1 The semi-quantitative amounts of arsenic species found in fish are summarized in Table 5.6. The most notable feature in these speciation results is the presence of arsenobetaine in all fish species and individuals. Arsenobetaine is the major arsenic compound found in most marine fish16'27. However, none (i.e., less than 5 ppb arsenobetaine) was found in a previous study of freshwater fish including bass, pike, carp, yellow perch, striped perch, pickerel (walleye) and whitefish, even though methods that had been developed specifically to identify arsenobetaine were employed 2 2. Another remarkable result is the presence of arsenosugar X I in suckers. Arsenosugars have been found in only one other fish, a marine fish (silver drummer) that feeds on algae 2 8. Most of the arsenic in the silver drummer was as T M A O (rather than arsenobetaine) with arsenosugars X I and XII I in the muscle, as well as arsenosugars X , XT, X I I and XII I (with the structures shown in Table 1.1, Chapter 1) in the digestive tract. The source of the arsenosugars in the silver drummer fish was postulated to be its food source, brown and red algae. Suckers are bottom feeders and may also acquire arsenosugar X I through their food, which implies that some benthic organisms are able to synthesize arsenosugars. The results in the previous chapter imply that cyanobacteria or other bacteria are able to synthesize arsenosugars X and X I and hence it is not unreasonable to suggest that organisms in the sediment may be capable of doing so. The major arsenic compound in pike is D M A . Pike is a pelagic carnivorous feeder, meaning that it derives its food from the water and not from the sediment (e.g., other fish). The stomach contents of pike sampled for this study consisted of invertebrates and small fish. Invertebrates may contain D M A , or arsenosugars. Arsenosugars may be broken down to D M A (observed in the human body 2 9 and in mice 3 0). For example, as described in the next section, mussels and snails contain arsenosugars X and X I , and D M A , among other arsenic species, 182 although D M A is not the major species. It might be expected that pike would accumulate more arsenobetaine, because its diet includes small fish, which probably contain arsenobetaine. However, pike appears to preferentially accumulate D M A . High levels of D M A have been observed in other fish; mackerel contained up to 25% of its total arsenic as D M A 1 6 . Two types of unknown compounds are present in these samples. Unknown X in Table 5.6 was observed in very low levels in whitefish and walleye, and had an early retention time on the cation exchange system. It co-eluted with arsenate and arsenosugar X I , but attempts to identify the unknown by using other chromatographic systems were unsuccessful. Unknown Y compounds (in Table 5.6) appeared as two late eluting peaks on the ion-pairing chromatographic system as shown in Figure 5.2 (retention times approximately 700 s and 1000 s). These compounds did not co-chromatograph with arsenosugars X , X I , X I I or XIII in kelp extract, as illustrated in Figure 5.2. The chromatograms in Figure 5.2 are scaled individually to facilitate comparison of peak retention times. It was surmised that the compounds should be anionic i f they are late eluting on the ion-pairing system used, but they did not appear on the anion exchange system. Possibly they were irreversibly bound to the anion exchange column. Unknown compounds eluting at similar retention times (with respect to D M A ) on the same chromatographic system have been observed as arsenosugar metabolites in human urine 2 9. 183 3.5e+4 o ( 0 k_ X CO a. cu i2 tz o o Pike 2 PD Kelp extract 0.0e+0 4e+3 Q in o o h Oe+0 240 480 720 time (s) 960 1200 Figure 5.2. Chromatogram of protease digest (PD) of a Yellowknife fish (Pike 2), showing small amounts of two unknown compounds ( U K - Y ) mentioned in Table 5.6. Ion-pairing chromatography (CI8 column, l O m M TEAH /4 . 5 m M malonic acid, p H 6.8, 0 . 1 % M e O H ; see Table 5.2) with I C P - M S detection was used. Abbreviations for arsenic species are in Table 1.1. 184 Branch et al. observed that the trypsin digest of a single specimen of plaice contained more D M A than the organic extract, and they postulated that partial degradation of arsenobetaine to D M A by trypsin was taking place. To determine i f the presence of D M A in pike, and the unusual presence of arsenosugar in sucker was a result of the protease digestion, rather than representative of the arsenic content in the fish, MeOH/water (1:1) extractions were performed for comparison. N o notable differences in arsenobetaine content in sucker are seen between the protease digestion results and the MeOH/water extraction (Table 5.6), although a more significant difference is observed in the amount of D M A extracted. The greatest difference appears for sucker 2, where not only is more D M A present, but less arsenosugar X I is observed when MeOH/water was used. The reasons for this are not clear. Possibly the arsenosugar in this matrix is bound to protein and the protease digestion is required to release it. Likewise, some D M A may be bound to a product of the protease digestion that precipitates or binds irreversibly to the chromatographic columns, whereas it may be effectively extracted as a free ion by using MeOH/water and can then be detected. Another possibility is that arsenosugar X I breaks down to form D M A during the MeOH/water extraction process. For pike, no significant differences are seen in arsenobetaine amounts by using the two methods (Table 5.6). Less D M A is extracted by using MeOH/water for both fish, which is the opposite observation as that for the sucker. Less unknown compound appears after MeOH/water extraction compared with protease digestion as well. Some of these compounds were probably released by the action of the protease on the matrix, and were probably not available for extraction by MeOH/water. When protease digestion was used, the sum of arsenic species observed (Table 5.6) is greater than 100% of the total arsenic (Table 5.5) for pike 1; however, considering that standard deviations are at least 30%, the amount detected after 185 protease digestion was within this amount and hence probably not significantly different from 100%. Extraction or digestion efficiencies were determined by using the relation: Extraction/digestion efficiency (%) = (sum of A s species)/(total As) x 100%. These values are somewhat higher for MeOH/water extraction compared to protease digestion for sucker, and lower for pike. Reasons for the non-extraction of certain species (i.e., greater amounts of D M A extracted from sucker, and smaller amounts of D M A extracted from pike, when using MeOH/water) are suggested above and these probably account for these differences in the sum of arsenic species detected. However, this comparison study is too small to draw any conclusions about the effectiveness of the different extraction techniques, except that the extraction of arsenic species appears to be matrix dependent and varies between different species of fish. Extraction/digestion efficiencies for most of the fish are lower than 100%, ranging from 19 to 78%o (except pike 1 by PD) . Protease digestion was expected to have solubilized most of the arsenic, although time constraints did not allow determinations of the total arsenic in the protease digestions to be made. However, i f most of the arsenic was solubilized, a large amount of arsenic is unaccounted for in these results. This arsenic may be bound to molecules such as peptides or incompletely hydrolyzed proteins, that may precipitate or remain irreversibly bound on the chromatographic systems used here. Arsenic is known to bind to cytosolic proteins from rabbit and rat l iver 3 1 ' 3 2 . In a previous study, 13.6 to 68% of arsenic remained in the residue from the trypsin digest for marine fish16. Some of the arsenic in the present study may have remained in the 186 residue after protease digestion, especially i f it was lipid bound. Lip id bound arsenic was postulated to be significant in mackerel, which is a fatty fish16. The lowest digestion efficiency observed in the present results was for whitefish 3, which is a fatty fish as well, as evidenced by an apparent lipid layer being present after the protease digestion procedure. Hence lipid bound arsenic may be significant for these samples, and experiments using lipase, to hydrolyze lipids, would be useful. To summarize, this study has shown that arsenobetaine is a major extractable arsenic species in freshwater fish. Arsenosugar X I was observed for the first time in freshwater fish, and D M A was observed to be the major arsenic species in pike. Although D M A has been linked to increased tumor growth in mice exposed to carcinogens3 3, the levels of D M A in these fish are too low to be of toxicological concern. Protease digestion and MeOH/water extractions gave comparable results in terms of arsenic species observed, although quantities varied, and more experiments are necessary to verify the differences. Extraction/digestion efficiencies were generally low, and again, more experiments are necessary to determine the reasons for this. 5.3.3. Freshwater shellfish 5.3.3.1. Total arsenic in shellfish The arsenic concentrations in freshwater mussels and snails collected from Yellowknife and from Campbell River are shown in Table 5.7. The freshwater mussel collected from Yellowknife was identified as Anadonta grandis simpsoniana and the snails are Stagnicola sp. The mussel from Campbell River is Margaritifera falcata. The mussels were collected from areas containing low levels of arsenic. The snails were collected from the marsh area at the Baker Creek outlet (location 1) and also from Baker Creek outside the mill area of Giant Mine (location 3). The mussel concentrations should thus reflect typical background concentrations, but the snails are most likely to have been impacted by high levels of arsenic. Table 5.7. Moisture content and arsenic concentration in freshwater shellfish. A l l shellfish are from Yellowknife, except where specified; C R = Campbell River; nd = not determined. Sample, sampling time (location) Moisture content (%)a (R=fresh weight/ dry weight) Arsenic concentration (ppm dry weight) Anadonta sp., June (16) nd 6.0 Anadonta sp., August (16) 90 (9.8) 7.4 Margaritifera sp. (CR) nd 3.1 Stagnicola sp., shelledb (1+3) 81 (5.2) 82 Stagnicola sp., whole b (1+3) 69 (3.2) 83 a Moisture content was calculated by using (R-l)/R*100%; R=fresh weight/ dry weight.0 nd = not determined. b The same species of snails was analyzed, but processing differed; see section 5.2.2. These concentrations obtained for arsenic in freshwater mussels (3.1-7.4 ppm dry weight) are similar to those for marine mussels, where values ranging from 8.47 to 15 ppm dry weight and 2.1 to 3.7 ppm fresh weight have been found 3 4 ' 3 5 ' 3 6 ' 3 7 . The levels of arsenic in these freshwater snails (Stagnicola sp.) are similar to levels found in marine gastropods, which can range from 4.2 to 233 ppm dry weight 3 8 ' 3 9 ' 4 0 . A freshwater snail contained 0.186 ppm (fresh weight) arsenic 4 1, which is an order of magnitude lower than the comparable concentrations in Stagnicola sp. in the present study, of 1.6 to 2.6 ppm arsenic (fresh weight). 5.3.3.2. Speciation of arsenic in shellfish The arsenic species that were determined following protease digestion (PD) and MeOH/water methods are shown in Table 5.8. 188 co £ c s CD o 5 • § 1 5 £ 3 co w O , t * <D ST g 3 .5 - .—. !=l <D ea ^ s e « O oc ,t-i ca O o <D ca .2? 8 CD 3 a a cu -5 (*> CD C ^ •9 § .3 -2 « ca c/3 £3 3 s u. i x: CD c/3 J-J 2 e <C o 5 ^ CD c/3 C L 6 n •a & <D . -c3 £ Vi I—J v-i H 3 CD U CJ ca (D t -6 0 CD 3 Q 5 O S O • - 3 2 > H «« s c CD II 3 ^ O e o U 00 I / } 3 H II (* U <D 3 8 CO CD II % co 73 . c 15 ca o fc- cD M a. O vi 1-3 .8 8 § S <U 3 M CO ca .a .3 Q O -1 i co Hi o < oo X X 3 CU T3 P on S ON co o c d V co O d V o d v d co d c/s 1« o co CN CN CN d CO d CN o d v s d v S i CO d co o d v CN d u g CO ( 3 s d v e C3 S Q CO CN o d v cu Z cu Z CN d ON d CN d V O d o d CO d CO CJ g °1 d S o d CN CN d CN o o d v cu Z a. Z co 2^ o d S u o g o o d v in o o CO o d cu Z cu Z VO d CN d d o d v o d v o d v d © CN — d ON CO CN CO CN o d V O oo o V O N d vo d o CN d 00 d oo vo oo CN CO CN O d v cu Z o d v CU Z o d v d d CN o d v t-; CO CO v d d s d v d s d v V O d o V CN <L> I 1 •S u ca & Si a) o I 8. .a V3 oo — 1 >w* 1 I a 1 •§ S « -8 NO CM u-i | | « Q 5 iS ca ca H a « o cu a cn ca a § © ft" "3 8 .3 C K > CD CD ^ •O & i o «J o II 5 Q ^ C/3 2 I s o •3 a B CD c j § u 189 The most striking result in Table 5.8 is the small amount, i f any, of arsenobetaine in freshwater mussels and snails. A small amount was extracted from the Campbell River mussel (3% of total arsenic extracted, fresh weight) but this was not reproducible when the extraction was repeated on a dry weight basis. A small amount (0.6%) is present in shelled snails. The absence of arsenobetaine has been observed for an estuarine mussel Corbicula japonica?4 and was suggested to be related to the low salinity of its environment. Glycinebetaine is used by marine animals for osmo-regulation 4 2, and it has been suggested that arsenobetaine is accumulated in marine animals because of its similarity to glycinebetaine. Therefore since estuarine and freshwater animals live in lower salinity regions, their need for osmo-regulation is reduced, resulting in less arsenobetaine entering the body of a freshwater animal and/or excretion of more arsenobetaine. These processes would result in less arsenobetaine being accumulated in the body of the animal. The absence of arsenobetaine in the animals analyzed in the present study supports the view that arsenobetaine is accumulated from the environment, i.e. by marine animals. Previous studies have also indicated that the presence of arsenobetaine in marine mussels is a result of bioaccumulation (i.e., from food) 4 3 . The major arsenic compounds in the mussels are arsenosugars X and X I (Table 5.8). The arsenic speciation in Yellowknife mussels collected in June and August differs only in the smaller amount of As (V) and a significant amount of an unknown species detected using the ion pairing chromatographic method, in the mussel collected in August (following PD) . The unknown arsenic species had a retention time on the ion-pairing system similar to that observed for one of the unknown compounds in pike protease digests (780 s, cf. 700 s in pike). As (V) is present in all mussel samples. MeOH/water (1:1) extracted As (III) from the Yellowknife mussel collected in August and the Campbell River mussel. In these specimens, the levels of inorganic arsenic found are not of toxicological concern to humans (should the mussels 190 be ingested). However, inorganic arsenic levels may be significant in situations where exposure of mussels to arsenic is much higher, and i f accumulation of arsenic by mussels takes place. Some MeOH/water extractions were performed to validate the arsenic species detected after using the P D technique, as for the fish samples. The sum of arsenic species is slightly higher by using MeOH/water extraction for the Yellowknife mussel collected in August (2.7 ppm cf. 2.2 ppm for PD) , but a lower sum results from MeOH/water extraction of shelled snails (23 ppm cf. 32 ppm for PD) . L ike the results for suckers, MeOH/water apparently extracts (or renders detectable) more D M A from Yellowknife mussels (1 ppm) than does P D (0.12 ppm). The reasons suggested for the differences in the sucker results probably apply here as well. More arsenosugar X is observed in protease digests, which may suggest that arsenosugar X I is being enzymatically degraded to arsenosugar X . The decomposition of arsenosugar X I to arsenosugar X in aqueous extracts during storage has been documented previously 3 4. However, this observation was not made for arsenosugar X I in sucker and hence it appears that individual matrices affect these processes. The results for the Campbell River mussel suggest that differences between fresh weight and dry weight extraction may be significant, in terms of extraction efficiency and species extracted or detected. Better extraction efficiencies have been observed for fresh weight compared to dry weight extractions for marine mussels4 4. For the two analyses of Campbell River mussels, differences in arsenic species are seen. Arsenosugars were observed in all samples, but small amounts of arsenite, M M A and D M A are observed following fresh weight P D of Margaritifera sp., whereas arsenite, arsenate and an unknown species are present following dry weight extraction (Table 5.8). The differences in arsenic species may be due to differences in the preparation technique (fresh weight or dry weight, and P D or MeOH/water extraction), or to 191 sample inhomogeneity. Sample inhomogeneity was observed by Shibata et ai. for a freeze-dried reference material and was considered to be quite significant. In this study, arsenosugar XIII was observed for a single fresh weight P D of the Campbell River mussel (results not shown), but it was not found in a duplicate fresh weight P D , nor in the dry weight MeOH/water extraction. The presence of arsenosugar XIII in the freshwater environment, had it been reproducible, would have been very significant, since up to now, only arsenosugars X and X I have been observed in the terrestrial environment as discussed in section 4.3.3.1 (Chapter 4). In snails (Stagnicola sp), the major extractable arsenic compound, in a proportion of 40-60% of arsenic extracted, is tetramethylarsonium ion. Inorganic species (As (III) and As (V)) amount to 25-40% of the total arsenic extracted, which is a major difference from marine snails. In a marine snail, Tectuspyramis, the major compounds were found to be arsenobetaine (35-67% of arsenic extracted) and tetramethylarsonium ion (6-26% of arsenic extracted)3 8. In other marine gastropods, arsenobetaine was also found to be the major arsenic compound, with inorganic arsenic, D M A , arsenocholine, the tetramethylarsonium ion or arsenosugar X as minor constituents 2 7 ' 4 0 ' 4 5. Uncharacterized trimethylarsenic (62% of total arsenic) and dimethylarsenic (27% of total arsenic) species were postulated to be present in a freshwater snail 4 1. The role of tetramethylarsonium ion is not clear. It has been suggested that it may serve as a precursor to or a decomposition product of arsenobetaine. However, its presence in snails, where the uptake or synthesis of arsenobetaine appears not to be significant, indicates that this is probably not the case. The tetramethylarsonium ion may arise from a metabolic pathway independent of one that would produce arsenobetaine, and is probably prevalent in freshwater gastropods. Methylation, possibly following the Challenger mechanism, might be taking place. The presence of M M A , D M A and T M A O in the snails, which would be intermediate compounds in the mechanism, supports this hypothesis. 192 Arsenobetaine is present in shelled snails in a small amount (0.6%), so the processes that occur in marine systems may also be taking place here, but to a smaller extent. Arsenosugar X was also detected in snails as a minor component (3.5%). The different sample preparation techniques used for the shelled snails may indicate that protease digestion releases more arsenic than MeOH/water extraction, although more studies should be done to verify this. The MeOH/water extract of snails was incompletely analyzed (by anion exchange and ion-pairing chromatography only), but the total amount of arsenic given in Table 5.8 reflects that which would be seen i f cation exchange chromatography was used, within the estimated uncertainty of 30%. Extraction/digestion efficiencies are lower than 50% for all samples analyzed. The same reasons as those given for the fish extraction efficiencies apply here: protease digestion and MeOH/water extraction may not solubilize all the arsenic, or the solubilized arsenic may not be detectable by using these chromatographic systems. When MeOH/water (1:1) was used to extract arsenic from marine mussels (Mytulis edulis in all cases) in other studies, only 52 to 60% of arsenic was extracted 3 4 ' 3 6. In another study, 47% of arsenic in Mytulis edulis was accounted for by H P L C - I C P - M S analysis as water soluble species, with 17% extracted in chloroform, and 32% remaining in the solid residue 3 5. Arsenic bound to lipids or other types of molecules such as proteins appears to be significant in mussel matrix. Another extraction method (MeOH/water, 9:1, with rehydration of dry weight samples before extraction) has proven to yield higher extraction efficiencies (95-100%) for marine mussels, although a significant amount remained unaccounted for after chromatographic analysis4 4. Chitin is a component of snails, since the radula (similar to a tongue) and parts of the digestive system are composed of this material 4 6. Protease digestion is inadequate for the solubilization of chitin and more specific enzymes such as chitinase and |3-(l-3)-glucanase are 193 necessary ' . In other studies, extraction of the marine snail T. pyramis with methanol was unable to dissolve 12 to 25% of total arsenic 3 8 which are significant amounts, but not as great as those unaccounted for in the present study. Mollusk shells, made of calcium carbonate crystals within a protein framework known as conchiolin, were found to require rigorous extraction with 2 M H Q 4 9 to dissolve arsenic. Hence it is expected that the mild sample preparation used in this study would not be capable of extracting arsenic from shells. This reasoning explains the observation that the extraction efficiency of whole snails is about half that of shelled snails, since the sample was probably diluted by the presence of shells from which arsenic could not be extracted. Arsenic is most likely present in snail shells, because it has been found in other mollusk shells 4 9. To summarize, the speciation of extractable arsenic in freshwater shellfish, containing very little or undetectable amounts of arsenobetaine, supports the suggestion that arsenobetaine accumulation in marine animals is related to osmo-regulation. Arsenosugars X and X I are the major compounds in freshwater mussels, similar to findings in marine mussels. Tetramethylarsonium ion is the major compound in freshwater snails, although inorganic arsenic is present in significant levels as well. The high accumulation of inorganic compounds in snails could present toxicological problems for higher trophic organisms. Large amounts of arsenic remain unidentified and this could be lipid or chitin bound. More work is necessary to elucidate the nature of this arsenic. 5.3.4. Plants Higher plants were collected from the Yellowknife area to determine the speciation of arsenic in plants. Sampling areas (refer to Figures 5.1a and 5. lb) included Kam Lake which 194 received accidental mining effluent discharges in the early 1970s from the Con Mine, and has also been influenced by raw sewage from Yellowknife 5 0 (location 12). Baker Creek, currently receiving mine effluent from the Giant Mine, was sampled in different areas: the marsh (location 1) and mill (location 3) areas, as well as an intermediate location in the creek (location 2). Niven Lake was also chosen as a sampling location, because it was a former sewage pond and was expected to contain residual elevated levels of arsenic and other metals, and because of prolific plant and algal growth. Two locations are specified, one on the east side of the lake, near the site of a former dump (location 10) and the other at the north end of the lake where the lake drains into Back Bay (location 11). Cattail, Typha latifolia, was common to all the sampling locations and horsetail, Equisetum fluviatile, was found in K a m Lake and Baker Creek. These two plants were sampled (shoots only) from different sites to determine i f any location specific differences in arsenic speciation could be seen. The aquatic plants, burweed, Sparganium augustifolium, and pondweed, Potomogetan richardsonii, were sampled from the water in Yellowknife Bay near a former Giant Mine tailings pond (location 6). 195 5.3.4.1. Total arsenic in plants The concentrations of total arsenic in plants are summarized in Table 5.9a. Uptake of arsenic by macrophytes was studied previously 4 for plants collected near Con Mine and some of these results are also shown for comparison in Table 5.9b. Table 5.9a. Arsenic concentrations in plants from Yellowknife. Sample, Time Location Arsenic concentration (ppm dry weight) Emergent plants (shoots) Horsetail Equisetumfluviatile, June 12 30 Horsetail Equisetum fluviatile, June 1 48 Horsetail Equisetum fluviatile, June 3 260 Cattail Typha latifolia, June 2 5.0 Cattail Typha latifolia, June 11 0.52 Cattail Typha latifolia, June 12 3.8 Bidens cernua, August 10 100 Submergent plants Milfoi l , Myriophyllum sp., June 11 78 (whole plant) Milfoi l , Myriophyllum sp., June 10 39 (whole plants) Milfoi l , Myriophyllum sp., August 10 17.4 (whole plant) Duckweed, Lemna minor, August 10 28 (whole plants) Burweed Sparganium augustifolium, 6 2.5 August (shoots) Pondweed, Potomogetan 6 20 richardsonii, August (shoots) 196 Table 5.9b. Arsenic concentrations in plants from Yellowknife, previous study Sample, Time Location Arsenic concentration (ppm dry weight) Emergent plants (shoots) Typha latifolia Equisetum fluviatile Triglochin palustre, arrow grass Submergent plants Potomogetan pectinatus (whole plants) Myriophyllum exalbescens (whole plants) Sparganium sp. (shoots) near Con Mine near Con Mine near Con Mine near Con Mine near Con Mine near Con Mine 17.2; range <1.0-38 34; range 5.5-91 40 1219; range 190-4990 143; range 30-255 28 Typha sp. contains the lowest levels of total arsenic, and this is especially obvious when the levels are compared with levels in other plants collected from the same sample location. For example, compare Typha sp. (3.8 ppm) with Equisetum sp. (30 ppm) at location 12 (Kam Lake); and Typha sp. (0.52 ppm) with Myriophyllum sp. (78 ppm) at location 11 (Niven Lake). In a previous study4, the average concentration of arsenic in Typha sp. shoots was found to be 17.2 ppm dry weight with a range of < 1.0 to 38 ppm dry weight, which is higher than the levels found in this study (0.52 to 5.0 ppm dry weight). However, the small sample size in this study precludes any direct comparison, except that Typha sp. shoots contained the lowest levels of total arsenic in the previous study as well as in the present one. Dushenko et al. postulated that the lower levels of arsenic in Typha sp., as well as its abundance at all sampling sites (observed in 197 the present study as well) indicates a greater tolerance to arsenic contamination by using mechanisms for the exclusion of arsenic4. Equisetum sp. contains high levels of arsenic at all the locations from which it was sampled (Table 5.9). Most likely the levels in these samples of Equisetum sp. are elevated (30 to 260 ppm dry weight) compared with background levels (5.5 ppm dry weight) in Equisetum sp. (sampled from Grace Lake, a location relatively unaffected by mine waste discharge) reported in the previous study4. The other emergent plant in the present study is Bidens cernua, a flowering plant that grew prolifically at the edges of Niven Lake in August, and the arsenic concentration in the shoots is 100 ppm dry weight. This concentration is higher than the average concentrations of arsenic found for all of the emergent plants sampled in the previous study (Table 5.9)4. The submergent plants Sparganium sp., Potomogetan richardsonii, Myriophyllum sp. and Lemna minor contain arsenic in concentrations ranging from 2.5 to 78 ppm dry weight. The previous study showed that submergent plants contained more arsenic than emergent plants4 (Table 5.9) but the sample size and number of sampling locations in the present study was too small to make a direct comparison. The concentration of arsenic in Myriophyllum sp. appears to decrease throughout the summer at the same location (39 ppm dry weight in June compared to 17.4 ppm dry weight in August), although this observation needs to be statistically verified. Grasses accumulate arsenic from soil at a faster rate when physiological growth of the plant is significant and accumulation of arsenic slows down when plant growth slows 5 1 . As described in Chapter 4, higher levels of arsenic were observed for fleabane and sedge at the end of the growing season. The results for Myriophyllum sp. indicate that an opposite trend is taking place, (i.e., arsenic concentration is lower at the end of the growing season) although the reasons for this are unclear. Possibly the rate of arsenic depuration exceeds that of uptake while plant growth is taking place. 198 5.3.4.2. Arsenic speciation in plants The arsenic species found in the plants collected in Yellowknife are listed in Table 5.10. The most noteworthy feature is the predominance of inorganic arsenic (As (III) and As (V)) as the extractable species, amounting to greater than 90% for all plants except for Lemna minor, which contains 80% of extractable arsenic as inorganic arsenic. For Equisetum sp. and Sparganium sp. inorganic arsenic forms were the only arsenic species extracted. 199 J 3 c at J 3 3 CD c3b CD ret o tu CJ c*> Q « M "Si O 43 00 °5 -a 3 O O X s "3 C H ^ 1 c/3 CD ' o C/l < C+H O S 3 C ' « Q 2 O CD M ex c / j ecj C + H 3 o X co c nj J 3 +n C/3 CA CD •a CD c n * C + H O c n +-» G 3 O J = Is H o o 2 « C + H O 3 .-a 3 42 I •a C N Tf I O co O C N — Tf r-VO rt M o V o V m - H o o © d V V t -co C N Ov o o d v o C N ~-> —< in C N m C N C N O m o Tf VO o „ r -o w Tt Tf VO t - ' C J o C N © C N O C N O _ C N © o 0> d V 0> © V 0> C N © © ' V C N © © V CN © © V C N © © V — cj ? s — u ? l - H © Tf © © V Tf © C N © © C O © V © V VO co © C N Tf C N X a C N u ~ a a 3 5 at a & 3 VO, ab 8 I C3 C 6 0 ^ I i f Si M 0, C I. ST I 5 ! © C N s I c •—> ts O H CO 5 a I It C O C N Tf vo C O © Tf © C N C N C N C N C N _ • — -© © © H H © © © © © © r - 00 © © ' © © ' © © © © © ' ©• o F - H V V V V V V V V V V © © ' C N d •n © C N © VO © 6 0 C o s: 53 c S cu VO C N © 9 0 © © V © V C J f ) © © © C N VO © © § Tf 00 C N C N Tf V V a © © ' © © ' © vo © / — ^ ^ — ^ © " / I © >n f-H C O d © © ' © © Tf ^ ' 1 — ' H H l - H C O VO © Tf Tf ^ — ' Tf © C N © C N IT) Tf Tf in vS" y—V © © © C N © H H © © C N vo © 1 — ' 00 © ' ^ — ' N -00 C N ~ — ' ^ ' © © ^—^ © * — ' VO in 00 C N co l - H •n C O © rH C O © r-~ C N vo © © ' © © C N rH C N © H H I cf a cu y co K c5 200 The proportions of arsenic species, especially A s (III) and A s (V) , are consistent between plants of the same species, as seen for Typha sp., Equisetum sp. and Myriophyllum sp. collected in June (see Table 5.11). The major arsenic species extracted from Typha sp. is A s (V), for Myriophyllum sp. (June) it is As (III), and equal amounts of A s (V) and As (III) were extracted from Equisetum sp. A s mentioned in Chapter 4, A s (III) is less toxic to periwinkle than As ( V ) 5 2 , and may be less toxic to higher plants from Meager Creek. Its presence in plants from Yellowknife may indicate that reduction of arsenate is taking place by the plant as a detoxification mechanism. The major species of arsenic in all water samples was arsenate (Table 5.3). However, the major extractable species from Baker Creek sediments was arsenite (see Table 5.3) which may be taken up unchanged by plants. Table 5.11. Percent arsenic species of total arsenic extracted from plants (SD) a . Plant As (III) As (V) Methyl Sugars M e 4 A s + ( D M A + ( X + XI ) _____ M M A ) Typha latifoliab (8) 4.8 (4.4) 0 0 Equisetum fluviatileh 50(10) 50 (10) 0 0 0 Sparganium augustifolium .30 70 0 0 0 Potomogetan richardsonii 11.8 78 7 1.6 5.9 2.0 Myriophyllum sp., June d 71 (12) 25 (14) 3.4(2.2) 0 0.1 (0.1) Myriophyllum sp., August 26 67 3 5.5 0.9 0 Lemna minor 17.3 63 7.2 10.7 1.7 Bidens cernua 8 88 3.4 0 0.5 Average for submergents 21(8) 70 (?) 3.6(3.3) 4.4 (5.0) 0.9(1.1) collected in August 6 a SD = standard deviation, calculated for plants of the same species sampled from different areas and with different absolute concentrations of arsenic. Where no SD is indicated, only one sample was collected, so n=l. b ForSD, n = 3. °ForSD, n = 2.. d Submergents collected in August include: Sparganium sp., Potomogetan sp., Myriophyllum sp. and Lemna minor, for SD, n = 4. 201 The difference between arsenate and arsenite distributions for Myriophyllum sp. between June and August is interesting to note. The sample collected in August contains the same proportion of arsenate as the June sample contains of arsenite (approximately 70%). This may indicate that oxidation is taking place throughout the summer, or it may indicate a slowing of the microbial activity (in the plant's environment) or plant activity to produce A s (III), which may be related to decreased plant growth in August. In general, submergent plants collected in August contain predominantly A s (V) (see Table 5.11) and therefore the trend observed for Myriophyllum sp. may extend to other submergent plants. Dushenko et al4, found that submergent plants accumulated more arsenic than emergent plants (Table 5.9). They postulated that leaf uptake from the surrounding water column was significant, and that no arsenic exclusion mechanism exists for submergents. The presence of arsenate as the major extractable arsenic compound in submergents after a summer of exposure may mirror the surrounding environment, and indeed, the major arsenic species in Yellowknife waters is arsenate. Minor amounts (less than 5%) of methylated arsenic ( M M A and D M A ) were extracted from all samples of Typha sp. (Table 5.10). The absence of methylated arsenic from extracts of other plants collected from the same location as Typha sp. (such as Equisetum sp. at Kam Lake) may indicate some specificity on the part of Typha sp. to accumulate these species or to methylate arsenic from its surroundings. Minor amounts of methyl arsenic are also observed in Potomogetan sp., Myriophyllum sp. collected in both June and August, and in Bidens cernua. A larger amount (7.2%) of methyl arsenic is present in Lemna minor. In all these species except Typha sp. and Equisetum sp., Me4As+ was observed at some point (e.g., in a.Myriophyllum sp. specimen in June, but not in August) Arsenosugars are also present in Potomogetan sp., 202 Myriophyllum sp. and Lemna minor. Evidence exists for both the synthesis and uptake of methyl arsenic (as M M A and D M A ) from its environment, as discussed in Chapter 4 5 2 > 5 3> 5 4 Me4As+ was found in a vascular plant from Meager Creek and its presence in these samples, albeit at very low levels, indicates a wider distribution than might otherwise be expected, considering the rarity of its presence in sediments or waters, as discussed in Chapter 4 5 5 - 5 6 . Halophytes apparently contained T M A O , M e 4 A s + , arsenocholine and arsenobetaine57 and the presence of these compounds was attributed to the presence of similar compounds in the surrounding water. The discovery of arsenosugars in some of these plants (Table 5.10), amounting to as much as 11% of extracted arsenic (absolute concentration 0.68 ppm dry weight) in Lemna minor, represents the first finding of arsenosugars in higher terrestrial plants. Halophytes (salt marsh plants) were proposed to contain an arsenosugar but no identification (i.e., comparison with standard arsenosugar compounds) was carried out 5 7. Interestingly, only arsenosugars X and X I are observed in the present study, which has also been the case so far for Meager Creek samples, and Yellowknife fish and shellfish. The arsenosugars are found only in submergent plants. Myriophyllum sp. and Lemna minor were growing in physical contact with algae, which may be a possible source of arsenosugars (mentioned in the next section). Submergent plants, in general, contain a larger proportion of organoarsenic species (on average, 9% of extracted arsenic, and up to 20% of extracted arsenic for Lemna minor, see Table 5.11) compared with emergent plants (up to 4.8 %). The reasons for this trend are unclear. Due to their propensity for uptake from their environment, as discussed by Dushenko et al.4, submergent plants may be acquiring organoarsenic from their environment, but the possibility of synthesis by the plants cannot be discounted. Extraction efficiencies are less than 100% in most cases, although two cases of efficiencies greater than 100% were obtained. In both these cases (Equisetum sp. from Baker 203 Creek marsh and Typha sp. from Niven Lake) the standard deviations in the speciation results are about 30% and the error in these numbers may account for these unusual results. The lowest extraction efficiencies are observed for Potomogetan richardsonii, Bidens cernua and Lemna minor. Unextracted arsenic could be bound to lipids, or to cell wall components, including insoluble cellulose, calcium or magnesium pectates, or lignin. 5.3.5. Algae and microbial mats Algae and microbial mats were sampled from a few locations in Yellowknife. Microbial mats were sampled from standing water next to, but not a part of, Baker Creek (location 4, Figure 5.1a), in both June and August. In June only the microbial mat was present in the standing water, whereas in August other plants (Typha sp. and Equisetum sp.) were present, but very little of the microbial mat was present. A microbial mat was also sampled from a small pond near a former Giant Mine tailings pond (location 5). Location 10 in Niven Lake was sampled for two different types of algae, which were painstakingly separated from each other and other organisms, and washed thoroughly. The exact composition of the microbial mats from locations 4 and 5 was unknown and most likely included a combination of bacteria (including cyanobacteria) and freshwater algae (including diatoms). The organisms possibly belong to genera such as Microcystis, Oscillatoria, Spirulina or Aphanizomenon, which have been found in other freshwater microbial mats 5 8. Two species of green algae were sampled from Niven Lake: Enteromorpha intestinalis, and Algae 1, of unknown identification, but possibly Cladorpha sp. 204 5.3.5.1. Total arsenic in algae Arsenic concentrations in the microbial mats are high, ranging from 390 to 2500 ppm dry weight (Table 5.12). These levels are higher than those found in Meager Creek microbial mats (maximum of 278 ppm dry weight). Table 5.12. Arsenic concentrations in algae from Yellowknife. Sample, Time Location Arsenic concentration (ppm dry weight) Microbial mat, June 4 2500 Microbial mat, August 4 1100 Microbial mat, June 5 390 Enteromorpha intestinalis, August 10 6.6 Algae 1, August 10 30 A bioconcentration factor (BCF) can be calculated for these algae, by using fresh weight concentrations and the following relations, where fw is fresh weight and dw is dry weight: [Asftv] = [Asdw]/R; where R = fresh weight mass of sample/dry weight mass of sample B C F = [ASfwMASwater] Factors of 0.86 and 0.38 are calculated for the microbial mats sampled from location 4 in June and August, respectively (R = 6). I f the factors are less than 1, then no bioconcentration is taking place 4 and this is likely the case for these samples. 205 Concentration factors can be calculated for the samples from location 10 as well; they are 9 (R = 10) for Enteromorpha intestinalis, and 68 (R = 6.5) for Algae 1. The water concentrations used for B C F calculations for the August samples are those determined in the June samples in Table 5.3 because concentrations of arsenic are not expected to change by more than about 50% in a location, from June to August 6 0 . The B C F s obtained indicate that Enteromorpha intestinalis and Algae 1 bioconcentrate and bioaccumulate arsenic. B C F s for marine and freshwater algae have been reported to be as high as 200 to 3000 6 1 . 5.3.5.2. Arsenic speciation in algae The arsenic species in the microbial mats sampled from location 4 in June and August show very few differences (See Table 5.13). The major species in these samples are inorganic arsenic, and the proportion of arsenite appears to increase slightly from June to August, where [arsenite]/[arsenate] was 0.47 in June and 0.70 in August. On the other hand, the proportions of D M A and arsenosugar X , although small in June, decrease to trace levels in August. The reasons for this are unclear and the differences are probably too small to draw conclusions. 206 s-< a r — I CD «n > .2? S PH 3 H - C cD c/3 O VH Cd. J- * a <D B 3 3 - o S « \E1 c/3 C < " 8 o !>-> • ° CO CD —: a Q ° cX Q ^ co S " ' C/3 - — - C/3 a ^ CD ' - ^ * 3 5 | > o o 8 aa «> CD T3 - s 2 •*H C/3 C2 CD B '3 0> CD C H C/3 ^ .a ^ g g C/3 C/3 a o •a cd a c CD a a o U • t-i Id cd i - J £ £ 1 CD +H o 0 0 0 0 3 0 0 X cd oo 3 0 0 _«J 0 0 o O H on vo CN CN o d v CN o d v co - — ' 0 0 cd s "3 IE o VH o O N C O CN O d v cu O N % O 3 S cj <^  cd o b CN o d V co CN N O d o N O CU oo s. 3 cd a "3 s o i s oo -d-CO Tf d O N d N O d CN CN o co CN o d v CU 3 3 cd = 2 o S r-o d N O o o CN o d o d v O N d T t CN o d v T t N O C O d I T ) d IT) CN o CN d CN d C O d co d o d v 8-2 I I 0 0 3 < <3 SC 0 0 3 < c u OC 5? O 'cu O 0 0 ~ cd * C J X CU CN l/-> CU I s p § cu «3 ,0 >. H .a J3 cu g> & 1 * 5 " H I 2 o ° i CN £ 5 CU T 1 ci 8 >> CJ s cu '5 ia a o a b x tu # II oo 207 The microbial mat sampled from the pond at location 5 contains arsenate as its major species, and 12.6% of the arsenic extracted occurs as methylated arsenic and arsenosugars (Table 5.13). The absence of arsenite in this sample compared with samples from location 4 probably indicates a lack of organisms that reduce arsenate to arsenite. Arsenosugars X and X I (10% of arsenic extracted) are present in this sample, whereas only arsenosugar X (3.4% of arsenic extracted for the June sample) is present in samples from location 4. The finding of arsenosugars in these samples and in microbial mats from Meager Creek in levels ranging from 0.8 to 44% of water soluble arsenic (Chapter 4) indicates that their synthesis is not restricted to thermophilic organisms. That is, among the non-thermophilic organisms that make up the microbial mats from Yellowknife, some appear to be capable of synthesizing arsenosugars X and X I . The algae sampled from location 10 belong to the phyllum of Chlorophyta. Because they are macroscopic it was possible to thoroughly wash them without disrupting or losing the organisms. From Table 5.13, it can be seen that arsenate is the major water soluble arsenic species (90% for Enteromorpha intestinalis and 33% for algae 1) in these algae. Arsenate is the only inorganic species present, indicating that no reduction to arsenite takes place by, or in the environment of these algae. Appreciable amounts of arsenosugars are present, especially in Algae 1 (sum of arsenosugars X and X I was 59% of extracted arsenic). This represents the highest proportion of arsenosugars found in freshwater organisms from both the Yellowknife and the Meager Creek environment. The terrestrial cyanobacterium Nostoc sp. contains the highest proportion of arsenosugars in any terrestrial organism (arsenosugar X was the major arsenic compound extracted)6. The presence of only arsenosugars X and X I (and not arsenosugars X I I and XIII) in the present study are in agreement with the findings from Meager Creek. These freshwater algae are probably synthesizing arsenosugars, although the possibility of uptake from microscopic organisms living in close physical contact with the algae and possibly 208 excreting arsenosugars cannot be discounted based on these studies. A unicellular marine diatom is apparently capable of synthesizing arsenosugars62, and freshwater analogues of these microscopic organisms may exist. L o w extraction efficiencies are observed for algae and especially for microbial mats (Table 5.13). L o w extraction efficiencies were also found for Meager Creek microbial mats (3-33%) and algae (1.1-7%). Only 1% of total arsenic was extracted from the microbial mat from location 4 that contained the highest levels of arsenic of all algae samples (2500 ppm dry weight). The low extraction efficiencies from microbial mats might be a result of arsenate coprecipitatation with iron and manganese oxides onto or in the microbial mats, making the arsenic insoluble in MeOH/water (1:1). A s discussed in Chapter 4, evidence exists to suggest that biomineralization of arsenic in microbial mats could take place 6 3 ' 6 4 ' 6 5 . The highest extraction efficiencies are observed for Enteromorpha intestinalis (49%) and Algae 1 (20%). Lip id soluble arsenic, which would have been extracted to only a very small extent by using MeOH/water (1:1) may make up some of the unextracted arsenic in these algae samples. Arsenic may also be bound to cell wall components such as those mentioned in 5.3.4.2. 5.3.6. Mosses 5.3.6.1 Total arsenic in mosses The total arsenic concentrations found in mosses collected from Yellowknife are summarized in Table 5.14. Two samples of Drepanocladus sp. were found growing underwater (locations 1 and 4) and another sample of Drepanocladus sp. as well as the other mosses were terrestrial (see Table 5.14 for location numbers). The identity of mosses 1 and 2 are not certain 209 but they are most likely the same species, Fumaria hygrometrica. Although Drepanocladus sp. was identified by genus only, all specimens of this moss are of the same unknown species6 6. Table 5.14. Arsenic concentrations in mosses from Yellowknife. Sample, Time Location Arsenic concentration (ppm dry weight) Drepanocladus sp., June 1 1220 Drepanocladus sp., August 1 490 Drepanocladus sp., August 4 880 Moss l a , June 5 1130 Moss 2 a, August 7 1900 Pohlia sp., August 15 1310 Drepanocladus sp., August 15 770 Moss 1 and Moss 2 are suggested to be the same species, namely Fumaria hygrometrica. The terrestrial mosses contain, in general, the highest levels of arsenic, and the sample from location 7 (Giant Mine tailings pond) contains the highest amount (1900 ppm dry weight, see Table 5.14). For terrestrial mosses, washing was probably not adequate to remove the soil in which they were growing, which may explain the very high levels of arsenic found in them. It was possible to sample aquatic Drepanocladus sp. so that sediment was not included and thus the levels in these moss samples more accurately represent the actual concentrations of arsenic in the moss. Drepanocladus sp. from location 1 appears to decrease in arsenic concentration from June to August, although the sampling locations were not exactly the same and replicate samples were not taken to verify this observation. Meager Creek samples of Fumaria hygrometrica also showed seasonal differences, where the concentration of arsenic in a November sample was 210 lower than that of a July sample. As with the Meager Creek samples, the seasonal differences in these samples may reflect differences in arsenic uptake rates and accumulation. 5.3.6.2. Arsenic speciation in mosses The arsenic species found in mosses are summarized in Table 5.15. Inorganic species of arsenic are the major and only species extracted from all moss samples, except from Drepanocladus sp. From all three locations, Drepanocladus sp. (1, 4 and 15) contain more extractable A s (V) than As (III), whereas mosses 1 and 2, presumed to be the same species, show no trend. The most remarkable feature of these results is the occurrence of M e 4 A s + in Drepanocladus sp. sampled from location 1 in June and August, but the lack of it in the same species of moss sampled from the two different locations 4 and 15. The major arsenic species present in snails, which were living in the moss, is M e 4 A s + . In a similar way, arsenosugars are present in Fumaria sp. from Meager Creek (Chapter 4), but not in the same or similar moss species from Yellowknife locations. These results may indicate that the arsenic species extracted from moss reflect the surrounding environment. This supports the hypothesis that for mosses, the source of arsenic species is uptake from the environment, rather than de novo synthesis. The arsenic species found in mosses from Meager Creek and Yellowknife, compared with the arsenic species found in some organisms (snails in Yellowknife, microbial mats in Meager Creek) that live in close physical contact with the mosses, are shown in Figure 5.3. 211 3 O o> 4} "B. C N C N d v o m o o i o a CN CN o © v CN o d v _> a s O, j © s p 0 0 o o d v 8 d v 8 CO O 1 s Q < 0 0 3 < VO CN O N CN a 3 O CN cn oo 3 < O in in CN r~-oo 3 < •J: •2 •St © a. p co C N CO CO CO o o o O d d d d V V V V 0\ © d CN NO VO —' O o O VO d d d ON V V V d CN o 8 VO O vo O d d d d V V V V '—V o © •«—^ /• N . 1—1 O N NO CN , — 1 0 0 o CO © d •-—' 55(6) d 24.5 4.86 55(6) 10.8 1 CJ © s I 0 0 3 < © <U O oo — § * in a p a s s 3 "3 > » I-H y 2 a oo o. •S a CN on V3 c3 •a _s § 3 g ^ o « 3 •_ X > O c" o • -a 2 •> . 1) : " 3 o a 'o II s o •a u g w II ! Q i on II W 212 As(lll)+As(V) MMA+DMA um x+xi Me 4 As+ o __ C CO o c o o CU > CD c_ Y K moss Y K snails MC moss MC algae Figure 5.3. Relative amounts of arsenic species in moss and associated organisms from Yellowknife ( Y K ) and Meager Creek (MC) , showing similarity in speciation between the moss and the organism (snails or algae) living in close physical contact with it. Abbreviations for arsenic species are found in Table 1 . 1 . 213 Extraction efficiencies for moss samples are low, ranging from 1.7% for moss 2 from location 7 to 15% for Drepanocladus sp. sampled in August from location 1. L o w extraction efficiencies were observed for mosses from Meager Creek as well. Limited extraction might be expected from the terrestrial mosses from which soil could not be completely removed, since arsenic in soil occurs in mostly non-water soluble forms, such as minerals, or adsorbed onto iron and manganese oxides. Arsenic appears to be bound to other plant components of the mosses for the non-terrestrial species, which may include lipids and cell wall components (see section 5.3.4.2.) 5.3.7. Lichens and mushrooms Pixie cup (PC) lichens (Cladonia sp., not positively identified as the same species) were found at different locations: P C I near Kam Lake (location 12), P C 2 near M e g Lake (location 13), and PC3 in the Con tailings pond area (location 15). Other lichens were collected as well near M e g Lake (location 13) and one species was found on rocks close to the water line of the Giant Mine tailings pond (lichen 4, from location 7). Identifications are Cladina sp. for lichen 1 and Cladonia sp. for lichens 2 and 3 (lichens 2 and 3 were not the same species). The identity of lichen 4 remains unknown but it was not Cladonia sp. Lichens 1, 2 and 3 were growing together. Mushrooms Paxillus involutus, Psathyrella candolleana and Leccinum scabrum were found near M e g Lake (location 13). M e g Lake is the first lake in the Meg-Keg-Peg drainage system, into which Con Mine effluent flows, and therefore it is the first lake to receive Con Mine effluent. The shaggy mane mushroom, Coprinus comatus, was found growing in the dry and sandy former Giant Mine tailings pond (location 8). Puffballs Lycoperdon pyriforme were sampled at two different locations. Mature specimens were found at the edges of the currently used Giant Mine tailings pond (location 7), and younger specimens were found at the edges of the Con Mine tailings pond (location 15). 5.3.7.1. Total arsenic in lichens and mushrooms Total arsenic concentrations in lichens and mushrooms from Yellowknife are summarized in Table 5.16. Levels of arsenic in lichens are higher than the levels in Meager Creek samples, which contained a maximum of 4.79 ppm dry weight of arsenic (Chapter 4). The highest concentration of arsenic in Cladonia sp. is observed in the sample from the Con Mine tailings pond, location 15 (520 ppm); it was likely to have been submerged at times of slow drainage from the tailings pond. The highest level of arsenic in all lichens occurs in Lichen 4 from the Giant Mine tailings pond (location 7). This was also the sampling location for the puffball mushroom, Lycoperdon sp., which contains the highest level of arsenic among mushrooms. The lichens that were sampled together, Lichens 1, 2 and 3, contain similar levels of arsenic, with an average of 47 (SD of 9) ppm dry weight. This may indicate that these different species of lichens accumulate arsenic to a similar extent. 215 Table 5.16. Arsenic concentrations in lichens and fungi from Yellowknife. Sample, time Location Arsenic concentration (ppm dry weight) P C I , June 12 14.3 P C I , residue3, June 12 6.4 PC2 , August 13 29 PC2, residueb, Aug. 13 15.9 (0.7) c PC3, Aug. 15 520 Lichen 1, Aug. 15 38 Lichen 2, Aug . 15 49 Lichen 3, Aug. 15 55 Lichen 4, Aug. 7 2300 Paxillus involutus, Aug. 13 36 Psathyrella candolleana, Aug. 13 13.6 Leccinum scabrum, Aug. 13 8.3 Coprinus comatus, Aug. 8 410 Lycoperdon pyriforme, Aug. 7 1010 a Sample obtained when the residue remaining after MeOH/water extraction was acid digested. b Residues following duplicate extractions, as for PCI (location 12). 0 Standard deviation obtained from total arsenic content in residues of duplicate extractions. Some of the mushrooms contain higher levels of arsenic than mushrooms collected from uncontaminated areas. For example, the specimen of Lycoperdon sp. analyzed in the present study contains 1010 ppm dry weight of arsenic, whereas the published background values are 0.46 to 2.81 ppm dry weight for the same family of mushrooms 6 7. Lycoperdon sp. from the Con Mine tailings pond (location 15) was not analyzed for total arsenic because the sample size was sufficient only for speciation analysis. Total arsenic levels for Psyaihyrella sp. and Leccinum sp. collected from uncontaminated areas were less than 0.2 ppm dry weight 6 8 ' 6 9 and for Paxillus involutus, they were 5.7-5.9 ppm 6 8 . Arsenic levels in Paxillus involutus, Psathyrella 216 candolleana and Leccinum scabrum in the present study are apparently elevated compared with the levels published for similar species collected from uncontaminated areas. The levels in the present study, however, are still close to the range of background concentrations (non-detectable to 15 ppm dry weight of arsenic) for most mushrooms 5 8 ' 6 7. The Coprinus comatus mushrooms from location 8 are appreciably elevated in arsenic concentration with respect to a published literature amount of < 0.1 ppm for Coprinus micaceus61. Its arsenic concentration is also elevated even when compared with the exceptionally high background levels (up to 130 ppm dry weight) found in Laccaria sp. 7 0 5.3.7.2. Arsenic speciation in lichens andfungi The arsenic species extracted from lichens and mushrooms are summarized in Table 5.17. The major arsenic species in lichens are As (III) and As (V), with the sum of these two species making up 62 to 93 % of the total extracted arsenic for the lichen specimens analyzed. Arsenobetaine has been found for the first time in lichens, and it is present in all the lichens sampled. Chromatograms of P C I and P C 2 extracts, obtained from cation exchange H P L C - I C P - M S analysis, are shown in Figure 5.4. Arsenobetaine was identified in P C I by spiking the sample extract with standard arsenobetaine and demonstrating co-chromatography of the suspected component with the authentic material (Figure 5.4a). It was identified in P C 2 (and other lichen samples) by matching the retention time of the presumed arsenobetaine peak in the sample with that of the standard (Figure 5.4b). 217 c 5 8 I 18 CJ cu h 1 a a CU CO J 3 > 3 § •a I x E ii 8 M S ' " C/3 a « J <» 5 / 3 g < I ° C + H » i o cu a CO Q O x CO C/3 C 4J a co CL c/3 So M co '2 CD § o C/3 a O 1 CO o C o U ° £ S » H .3 . cu +5 3 2 H o a Q O a o CO T3 00 < 1 o < 3 X 60 3 00 e 0) o c3 O 00 O CN T t CO CN T-H © CO CN CN d © O o O © d NO d d d T t © V d V V V d V © © CN © d v C O d oo d CN d 00 d r -T t CO CN OS (—, s °. d s © vo © © v _ < 5 © vO d T t d vo CN © T t © VO © V vo VO CN CN CO I T ) l-H * — 'CN co U U U Cu OH CH vo d © S 3 © V ON CO © vo CN CO X CN CN © o © © © © H H d © © V © V V V CO CN CN CN © © © © © © © © © © V V V V V VO d T t © © V 00 CO IT) © CO d r -d p T t >o co CO - N T t . -. 00 © © © w T t vO CN vd T t T t CN co chen CO a <u .fl ^ O CO S C J JS o ^—s CO J —1 w —J ^—' T t CN </-> © T t s CJ J3 cj CJ CO •a i OJ CJ "3. co H> T 3 C S3 S a CO >> CO CJ X CJ S O CJ 60 cd §£• O \o B £ c 3 '-3 S o ^ <+H g s •= a S c o CO JO r-J • u ^ § •§•(2 1 60 ^ ">> 2 o -3 co '5 CJ E « s o o, g s »» s - C co a , CJ <j a N CH A ^ l CH § 2 S S 3 c3 <a a CJ •o u > 4> 2 •a a oo 8^-2 •i .2 JS > n ™ CJ a CJ •O 2 OH | w X a II z CO " O II W c W oo e« D 2 1 8 CD i5 IS I S 3 c o 3 o ion C-oo o co X P >< 2 2 n __3 o d v C O V O o d v CN d d 3 5 1! _, V O -"" •o 0 0 — - t S p 0 0 oo — < vd V O r-— « d r-o r-o X —-» X d © © V V d V © © — i © © d C O d S V V © © t— r— o © d © C O vo v O V V © © © .—> © © © o t © C N — * C O d © © © d V vo co V O r—< C N o © © C N C N d © © © © V V V V V — i V O cs V O d d d d co V O IT) O d vd — H — * C N C N © © C N © © © © s V V V O V CO d CN © -S e I is l l © s 5 s -a s © CN 0 0 r \ © co co I so §• CJ C N J^ 3 219 I I I I I 1 1 1 1 1 1 0 120 240 360 480 600 720 840 960 1080 1200 time (s) Figure 5.4. Arsenobetaine ( A B ) in lichens by using cation exchange H P L C - I C P - M S (see Table 5.2 for details). 5.4a. Chromatograms of Yellowknife lichen (PCI) extract and extract spiked with 50 ppb A B standard. 5.4b. Chromatograms of Yellowknife lichen (PC2) extract and 100 ppb A B and A C standards. See Table 1.1 for abbreviations, U K = unknown compound. 220 Arsenosugar X is present in minor amounts in two lichen samples, P C 2 (5% of extracted arsenic) and Lichen 3 (4% of extracted arsenic) from location 13. Arsenosugars were not detected in Lichens 1 and 2, which were growing together with Lichen 3 at location 13 (Table 5.17a). Some similarities exist for all lichen species that were sampled from the Con Mine Meg-K a m Lakes drainage system (locations 12, 13 and 15), summarized in Table 5.18. The proportion of A s (III) is similar for all samples (42-61% of extracted arsenic, see Table 5.18), but a broader range is observed for the relative amounts of As (V) in the lichens (12-44% of extracted arsenic). Amounts of arsenobetaine range from 4 to 11%, and similar amounts of T M A O were observed for the three lichen species (lichens 1, 2 and 3) that were growing together (4-6.5 % of extracted arsenic). Similarities in arsenic speciation between these three lichens might be expected, due to their common environment. The distribution of arsenic species found in the lichens collected near Con Mine (locations 12, 13, and 15) differs from that in the Giant Mine lichen (location 7) as summarized in Table 5.18. The major differences observed are the predominance of A s (V) extracted from Lichen 4, and the small proportion of arsenobetaine found in Lichen 4 (0.5%) compared with the amounts found in the other lichens (4 to 11%). These differences may be caused by differences in lichen species (and hence metabolism of arsenic), and/or differences in their environments. For example, the major form of arsenic is probably as arsenate adsorbed onto ferric hydroxide in the Giant Mine tailings and the major form of arsenic in the tailings pond water is arsenate60. Unlike lichen 4, the Con Mine lichen samples were less likely to be submerged in tailings pond water, except for one sample (PC3). The arsenic speciation might reflect the extent to which the lichen is directly impacted by the tailings water. 221 Table 5.18. Proportions of total arsenic extracted, in % of arsenic species for lichens and mushrooms. Sample (location) As (III) As (V) Methyl + Sugars A B T M A O + cations P C I (12) 44 44 1.9 11 0 P C 2 (13) 50 12 10 11 18 PC3 (15) 61 27 7.4 4.0 0 Lichen 1 (13) 55 26 8.6 5.1 5.9 Lichen 2(13) 60 18 5.1 10.3 6.5 Lichen 3 (13) 42 35 13 4.0 6.0 Average for Con Mine lichens (12, 13, 15) 52 (8) a 27(11) 7.7(3.9) 7.5 (3.5) 6.1 (6.5) Lichen 4 (7) 1.9 91 6.5 0.5 0 Lycoperdon sp. (7) 4.6 29 22 42 7.8 Lycoperdon sp. (15) 16 1.6 3.2 78 1.8 Standard deviation of proportions of each compound. The mushroom Lycoperdon sp. from the Giant Mine tailings pond (location 7) also contains a proportionally higher amount of arsenate, although the major arsenic species in both specimens of Lycoperdon sp. is arsenobetaine. Arsenobetaine was also observed to be the major arsenic species extracted from Lycoperdon echinatum (78%), Lycoperdon perlatum (88%) and Lycoperdon pyriforme (62%) in a previous study; minor components included As (V), As (III), M M A and D M A 6 7 . A n greater variety of arsenic species is observed in the Lycoperdon sp. from Giant Mine tailings pond (location 7) compared with that in Lycoperdon sp. from the Con Mine tailings pond (location 15); see Figure 5.4 and Table 5.18. The specimen from location 7 was observed to be in a more mature form, and this may account for the differences in speciation 222 i ] As(lll)+As(V) MMA+DMA AB | | AC+Me 4 As+ +UK 100 Figure 5.5. Relative amounts of arsenic species in the puffball mushroom Lycoperdon sp. from location 7 (Giant Mine tailings pond) and location 15 (Con Mine tailings pond). Differences in speciation are seen for Lycoperdon sp. from different locations. Abbreviations for arsenic compounds are in Table 1.1; U K = unknown compound. 223 observed. Again, the microbial environment influencing the fungus may also cause differences in arsenic speciation. The arsenic speciation has been determined for water-soluble species in the shaggy mane mushroom Coprinus comatus for the first time, and the major arsenic compound is arsenobetaine (88% of extracted arsenic). Minor components include inorganic arsenic, D M A , arsenocholine and an unknown arsenic compound. Paxillus involutus grew in abundance next to the Con Mine effluent stream. Its arsenic content is unusual because a major proportion of arsenic extracted (36%) is in an unidentified form (unknown compound Y ) . The peak for unknown Y was broad and it eluted near the retention time of arsenobetaine on the cation exchange system (see Figure 5.6 for chromatogram.) Neither unknown Y , nor a minor component, unknown X , could be identified as any of the arsenic compounds available to us as standards; some of the cationic arsenic standards are chromatographed in Figure 5.6. A n unknown peak possessing a retention time similar to that for unknown Y , on almost the identical chromatographic system, was observed in mussels by Larsen et al.35 The authors established that the compound was neither 2-dimethylarsinylethanol ( M e 2 A s ( 0 ) C H 2 C H 2 O H ) nor glycerylphosphoryl-arsenocholine ( M e 3 A s + C H 2 C H 2 0 - P 0 2 " - O C H 2 C H O H C H 2 O H ) . The other major arsenic compound extracted from Paxillus involutus is D M A (53% of extracted arsenic). As (III) and (V), M e 4 A s + , another unknown species (unknown X ) , and a small amount of arsenosugar X I (2.6% of arsenic extracted) occur as minor components. 224 2e+4 DMA Paxillus sp. extract 25 ppb standard mix 0 240 480 720 960 1200 time (s) Figure 5.6. Chromatogram of Paxillus involutus extract (diluted lOx), showing the presence of unknown compounds X and Y ( U K - X and U K - Y , respectively), as well as D M A , inorganic arsenic, and a small amount of Me4As+. Standards (including 25 ppb of M M A , A B , T M A O and A C ) do not co-chromatograph with U K - X and U K - Y . Abbreviations for arsenic compounds are found in Table 1.1. 225 The major arsenic compound found in Psaihyrella candolleana is arsenate, making up 63% of the total arsenic extracted. The next most abundant compound is arsenite, and minor components are M M A , D M A , arsenobetaine and M e 4 A s + . This mushroom species is classified in the same family as Coprinus sp. (Coprinaceae) 6 9. Similarities have been observed in arsenic contents of different mushroom species within the same family 6 7, and thus it is not surprising to find some of the same compounds (e.g., arsenobetaine) in Psathyrella candolleana and Coprinus comatus. Both mushrooms are edible but the major water-soluble arsenic species in Psathyrella candolleana, As (V) and As (III), are toxic to humans, whereas in Coprinus comatus the major species, A B , is not. Leccinum scabrum contains mostly D M A with minor amounts of inorganic arsenic. In comparison, the three major arsenic compounds extracted from Boletus edulis (Chapter 3) were arsenite, arsenate and D M A . These two mushrooms belong to the same family (Boletaceae) 6 9, and again, similarities in arsenic species can be seen. It has been suggested that in more "primitive" genera of mushrooms, arsenobetaine occurs much less frequently than in genera that are more highly evolved, such as puffballs6 7. On this basis, Paxillus involutus, and Leccinum scabrum may be more primitive mushrooms. The presence of arsenosugar X I in Paxillus involutus represents one of the first reports of arsenosugars in mushrooms. Arsenosugar X I was tentatively identified in an extract of Laccaria amethystina, but further chromatographic confirmation was considered to be necessary70. Small amounts (less than 1% of extracted arsenic) of arsenocholine were found in Coprinus comatus and Lycoperdon sp. Arsenocholine has been observed only in small amounts in marine samples, even though studies have demonstrated that this compound can be readily bio-transformed into arsenobetaine by sediments71. However, arsenocholine has been observed 226 previously in mushrooms, being one of the major extracted arsenic species in Amanita muscaria72 as well as in Sparassis crispa61. Other researchers 6 7 ' 7 3 have suggested that the fungus itself is responsible for synthesizing arsenicals, such as arsenobetaine, that had not previously been found in the terrestrial environment. The absence of these compounds in soil (although arsenobetaine was recently found in ant hill material 7 4) provides strong support for their hypothesis. However, experiments summarized in Chapter 3 indicate that synthesis of arsenobetaine or arsenocholine does not take place in pure culture of fungi. In the present study, the arsenic compounds found in a mushroom species (Lycoperdon sp.) appear to depend on the location, indicating that the surrounding environment has a strong influence on the extractable arsenic species from a mushroom. As well, it was shown in Chapter 3 that arsenobetaine in the culture medium was readily taken up by fungus. I f arsenobetaine was being synthesized by soil organisms, or organisms associated with the mycelia, it would be taken up efficiently by the fungus and not be detectable in the soil. Experiments with organisms cultured from the environment of cultivated or wild mushrooms might help to elucidate the source of complex arsenicals such as arsenobetaine, arsenocholine and arsenosugars. Extraction efficiencies for lichens and fungi range from 1.1% for Lichen 4 from location 7 to >80% for Paxillus involutus and Leccinum scabrum from location 13. The average extraction efficiency for lichens collected from the Con Mine area (locations 12, 13 and 15) is 21% with a standard deviation of 12. The lowest extraction efficiencies are observed for PC3 from location 15, and Lichen 4 from location 7, which are the lichens containing the highest levels of total arsenic. The mushrooms Coprinus comatus and Lycoperdon sp. contain the highest levels of total arsenic and their extraction efficiencies are the lowest. Extraction efficiencies for Lycoperdon sp. 227 in a previously published report ranged from 24% to 128% , whereas 68% of arsenic was extracted from Laccaria amthystina70 containing levels of arsenic comparable to those in Coprinus sp. and Lycoperdon sp. The present results suggest a negative correlation between extraction efficiency and arsenic concentration in lichens and mushrooms. However, the reasons for this apparent correlation remain unclear. A possibility is that some of the arsenic contributing to high total levels of arsenic is in a non-extractable form, such as in a mineralized form on the outside of the specimen, or bound to chitin or other cell components of the fungus. The distribution and metabolism of arsenic may not be uniform for different levels of arsenic uptake for fungi, which may explain differences in extraction efficiencies. Residues of two extracted lichens were analyzed for total arsenic (see Table 5.16) and extraction efficiencies from these results were calculated as: % Extraction efficiency = [(total arsenic)-(arsenic in residue)]/(total arsenic) x 100%. Based on this calculation, 55% of total arsenic was extracted from P C I , which differs from the extraction efficiency of 13% in Table 5.17 (calculated by summing the arsenic species detected). A n extraction efficiency of 45% was calculated by using the arsenic concentration in the P C 2 residue, which agrees quite well with the 42% reported in Table 5.17. The difference in extraction efficiencies for P C I indicates that some arsenic that was extracted was not observed by using the chromatographic systems available. This was suggested previously for other samples as well. 228 5.4. Summary A large amount of information about arsenic in the terrestrial environment was obtained from the study of Yellowknife biota. A n important observation was, as for the Meager Creek study, the low proportions of water soluble arsenic species (evidenced by low extraction efficiencies) in the majority of the biota studied. Arsenic that is not extracted is probably bound to lipids, cell components, proteins, or exists as minerals. Although the extractable portion can be approximated to represent the arsenic species that are bioavailable, the availability of the non-extractable arsenic cannot be assumed to be negligible, since the metabolism of such arsenic by higher trophic level organisms has not been reported. Freshwater fish were found to contain arsenobetaine, in contrast to previous reports. Sucker was found to contain arsenosugar X I , and probably obtains the arsenosugar from its food source, implying that benthic organisms are capable of synthesizing these arsenic compounds. The major extractable arsenic compound in pike was found to be D M A , but the source is unknown. Amounts of arsenobetaine in freshwater mussels were negligible or non-existent, supporting the idea that the uptake of arsenobetaine in the marine environment is related to osmo-regulatory processes. Mostly inorganic arsenic species were extracted from plants, mosses, algae and lichens. Plant species appear to be consistent in their uptake and metabolism of arsenic, which may indicate some transport mechanisms specific to the plant. Arsenosugars were found for the first time in higher plants. Freshwater green algae, as well as microbial mats, contain arsenosugars, probably indicating synthesis by the organisms. Arsenobetaine was found for the first time in lichens. Differences in arsenic speciation are seen for the mushroom Lycoperdon sp. from different locations, which may indicate that 229 complex arsenicals, such as arsenobetaine, are formed as a part of the fungus microbial community and taken up by the fungus, rather than synthesized by the fungus itself. References 1. Controlling Arsenic Releases to the Environment in the Northwest Territories: Discussion of Management Options; Environment Canada. Health Canada. G N W T Health and Social Services. Apri l 1997. 2. Bright, D . A . ; Coedy, B . ; Dushenko, W . T.; Reimer, K . J. Sci. Tot. 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Its compounds are used as flame retardants in plastics and textiles, as additives in metal alloys, as doping agents in semiconductors, and as antiparasitic drugs1. Because of the toxicity of some of its compounds, antimony is listed by the United States Environmental Protection Agency ( U S - E P A ) as a priority pollutant2. The determination of antimony species in the environment is necessary in order to assess the toxicity and mobility of antimony. Usually concentrations of naturally occurring antimony are about 5-10% those of arsenic3 and the two elements are often found together in mineral deposits4. Because elevated levels of arsenic are found in the Meager Creek and Yellowknife environments (Chapters 4 and 5), and because elevated antimony levels were found previously in biota from the Yellowknife environment5, we were interested in increasing the range of samples previously analyzed for antimony content. Although methods for antimony speciation by using H P L C with element specific detection have been developed 6 ' 7 ' 8, determination of methylantimony species other than Me3SbCl2 by using these methods have not been successful9. Therefore, we chose to use the method of hydride generation-gas chromatography ( H G - G C ) with A A S and M S detection for the analysis of antimony in environmental samples, in spite of the limitations and problems associated with this speciation method (see Chapter 2 and references at the end of this chapter 5 1 0 ' 1 1). 236 6.2. Experimental 6.2.1. Chemicals and reagents Antimony (V) and (III) standards were obtained as potassium hexahydroxyantimonate, K S b ( O H ) 6 (Aldrich), and potassium antimonyl tartrate, K 2 Sb2 (C 4 06H2 )2 (Aldrich). M e 3 S b C l 2 was synthesized as described elsewhere1 2. Stock solutions were made by dissolving these compounds in deionized water and diluting the resulting solutions to 1000 or 100 mg L" 1 as Sb. Standard working solutions were made by diluting the stock solution with deionized water as necessary. For hydride generation analysis, NaBFL, (reagent grade, Aldrich) was dissolved in deionized water fresh daily to provide a concentration of 2% w/v. Ammonium citrate buffer at a concentration of 0.05 M and p H 6 (1 M ammonium hydroxide, MicroSelect, Fluka, and analytical reagent grade citric acid, B D H ) and 1 M HC1 (Environmental grade, Alfa Aesar) were used for p H adjustment during derivatization. 6.2.2. Sampling and sample preparation Sampling was carried out at Meager Creek ( M C ) and in Yellowknife ( Y K ) , as described in Chapters 4 and 5, with sample locations shown in Figure 6. l a ( M C ) and Figure 6. l b ( Y K ) . Water was sampled by hand into polypropylene bottles that had been acid washed previously. Biota were sampled by hand, stored in Ziploc® bags and kept cool until processing. They were washed thoroughly with tap water to remove soil and other particles, rinsed with deionized (1 Mohm) water, and frozen. The samples were freeze-dried and pulverized to a fine powder for analysis. Snails from Yellowknife were dissected to remove the soft tissue prior to freezing. 237 Samples were digested with acid for the determination of total antimony content. The freeze-dried powders were accurately weighed (0.5 g ± 0.5 mg) into a 500 mL round bottomed flask (RBF). Concentrated nitric acid (3 mL, doubly distilled in quartz, Seastar, Sidney, B C ) and hydrogen peroxide (3 mL, 30% in water, reagent grade, Fisher) were added to each sample. The samples in the R B F s were boiled for 3 hours by using a heating mantle and a reflux apparatus13. After all the samples had cooled, the clear solutions remaining were diluted to 25 mL with deionized water and stored at 4°C until analysis. Extractions were carried out by weighing 0.5 g (± 0.5 mg) of the dried powders into 50 mL or 15 m L centrifuge tubes, adding 10-15 m L MeOH/water (1:1), sonicating for 20 minutes, centrifuging for 20 minutes, and decanting the liquid layer into a R B F . Each sample was sonicated and centrifuged a total of 5 times. The decanted extracts for each sample were pooled and rotovapped to near dryness (1-2 mL) and then diluted to 5 or 10 mL with deionized water. Moss (Drepanocladus sp.) from Yellowknife locations 1 and 4, and snails (Stagnicola sp.) from Yellowknife locations 1+3, were extracted in a larger quantity to permit analysis by using H G -G C - A A S and H G - G C - M S . Masses of 1 or 2 g were weighed out and extracts were made up to a final volume of 20 m L or 40 mL, respectively. 238 Figure 6.1a. Map (not to scale) of Meager Creek Hot Springs area showing sampling locations. 239 6.2.3. ICP-MS analysis for total arsenic and antimony concentrations Acid digested samples and water samples were analyzed by I C P - M S for total antimony content. A V G Plasmaquad PQ2 Turbo I C P - M S ( V G Elemental) outfitted with a peristaltic pump and injection loop for flow injection introduction was used. Parameters for the I C P - M S are given in Table 6.1. The m/z monitored were 121 (Sb), 123 (Sb) and 103 (Rh). Ac id digested samples were diluted with 1% nitric acid (doubly distilled in quartz, Seastar) and R h (10 ppb) was added as an internal standard to diluted samples and water samples. Quantification was carried out by using an external calibration curve derived from Sb standards made in 1% nitric acid and containing 10 ppb Rh. Table 6.1. Operation parameters for I C P - M S Feature Forward radio-frequency power Reflected power Cooling gas (Ar) flow rate Intermediate (auxiliary) gas (Ar) flow rate Nebulizer gas (Ar) flow rate Nebulizer type Quadrupole pressure Expansion pressure Parameter T350"w" <10 W 13.8 L/min 0.65 L/min 1.002 L/min de Galan 9 x 10"7 mbar 2.5 mbar 6.2.4. HG-GC-AAS analysis for antimony speciation The apparatus was composed of a semi-continuous flow, hydride generation system developed for arsenic analysis, 1 4 coupled to an atomic absorption spectrometer (Varian AA1275) fitted with an Sb lamp (Varian) operating at a wavelength of 217.6 nm, or at 231.4 nm for a few 241 samples (section 6.3.2). One modification was made to the basic apparatus in the form of using a gas-liquid separator1 5 that resulted in less analyte carryover. The apparatus consisted of Tygon tubing for the peristaltic pump, and P T F E tubing (1/8" OD) for the remainder. Data were collected from the A A S and processed directly by using an FfP 3390A integrator, or were analyzed with the aid of Shimadzu EZChrom software run on a P C . A peristaltic pump was used to deliver standard or sample solution (ranging from 5 u L to 200 p L for standards, and from 1 mL to 100 m L for samples) to mix with the acid or buffer and then to mix with a solution of NaBFL, (2% w/v) in a reaction coil. The gases evolved were separated in the gas-liquid separator and then swept by a flow of helium into a P T F E U-tube, where they were trapped at -196 °C. Continuous hydride generation and trapping were carried out for 3 minutes. The peristaltic pump was then stopped (making the system semi-continuous) and the U-tube was heated to 70 °C, allowing the gases to be swept with He at a flow rate of 40 mL/min onto a Poropak PS column, which was then heated from 70 °C to 150 °C at a rate of 30 °C/min, whereby the gases were separated. They were then detected by A A S . Semi-quantitative amounts were calculated by using external calibration curves. 6.2.5. HG-GC-MS analysis for confirmation of methylantimony species Extracts of samples and standard solutions of MesSbCb. were reacted with NaBFL, to form methylantimony hydrides. The reactions were performed in a 15 mL vial (sealed with a PTFE-faced silicone or neoprene septum, 16mm, Supelco) by using the appropriate volume of sample (5 m L for moss extracts and 10 m L for snail extract) or MesSbCb. standard solution, and 1 mL deionized water (for standards) and then injecting 0.5 mL of 6% NaBEL, solution through the septum. A l l reactions were carried out without the addition of acid or buffer, with the exception of one reaction in which M e 3 S b C l 2 standard solution was first made acidic by adding 242 an equal volume o f 1 M HC1, in order to generate M e S b H 2 and M e 2 S b H in addition to Me 3 Sb. These methods were qualitative only. For the analysis, a G C - M S system consisting of a Star 3400Cx gas chromatograph (Varian), equipped with a 1078 temperature programmable injector (Varian) and interfaced to a Saturn 4D ion-trap mass spectrometer (Varian) was employed. A gas tight syringe (1.0 mL, Gastight #1001, Hamilton) was rinsed with 5 mL of lab air and then used to inject 1 mL of headspace generated from the samples onto a capillary column (PTE™-5, 30m x 0.32mm, 0.25 um, Supelco 2-4143, poly (5% diphenyl / 95% dimethylsiloxane)). The injector was kept at 100 °C. The temperature program started at 40 °C, and stopped at 150 °C with a heating ramp of 15 °C/min. The parameters used are shown in Table 6.2. Table 6.2. G C - M S parameters. G C method Injector temperature Column temperature program Transfer line temperature Column M S method Mass range Scan time Segment length Ion Mode Multiplier Target Ionization current Manifold temperature 100 °C 40 °C, 15°C/min to 150 °C 200 °C PTE™-5 , 30 m x 0.32 mm, 0.25 115-180 m/z 0.4 s 4.5 min Electron Impact 2150 V 15 400 20 u A 260 °C 243 6.3. Results and Discussion 6.3.1. Antimony species and total antimony in environmental samples The semi-quantitative amounts of antimony species in environmental samples detected by using the method of H G - G C - A A S , as well as total antimony, determined by using I C P - M S , are shown in Table 6.3. The antimony contents in biota and water from Yellowknife and Meager Creek are summarized in Table 6.3a (biota) and Table 6.3b (water). Absolute detection limits of 1 ng for Sb (III) and methyl antimony species were estimated and relative standard deviations between replicate analyses can be estimated to be 20%. The biota samples were chosen for analysis by H G - G C - A A S based on their ability to meet one of two criteria: (a) the presence of antimony in sample extracts was observed during H P L C - I C P - M S analysis or (b) total antimony content greater than 10 ppm dry weight was measured. 6.3.1.1. Inorganic antimony species In all biota sample extracts and water samples, inorganic Sb (V) is the major antimony species (Tables 6.3a and 6.3b). Very few biota samples have been speciated for antimony previously 5 ' 1 6, but other researchers have also shown that Sb (V) is the major antimony compound in water samples 3 ' 8 ' 1 7 ' 1 8 ' 1 9 . Thermodynamically, Sb (V) is predicted to be the most stable oxidation state under most environmental, oxygenated conditions (pH 5 to 8 ) 2 0 and therefore it is not surprising to find that Sb (V) is the most abundant extractable species of antimony in these samples. 244 1 ) o S C/3 S H O O •a a is i — w w 5 "8 H -3 oo O CO 0 0 0 0 oo I o © r - H ° ° . 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"s a, 2 3 3 I—> u CU — i —> a j on on CN „ on cs — i CN + — * CN on on on -*• e S s a a a s o o o o o o o •a •3 •3 •3 •3 •3 •3 es u 53 U 8 3 cS o 3 o o o o  o -1 -1 - J —1 —1 246 Inorganic Sb (III) is present in Drepanocladus sp. (moss) samples from Yellowknife ( Y K ) location 1 and Yellowknife ( Y K ) location 4, as shown in Table 6.3a. Sb (III) is also present in all Meager Creek ( M C ) waters and water from Y K location 4 and location 11 (Table 6.3b). Other studies have shown that Sb (III) is produced in the photic zone of an estuarine inlet, which may indicate that biological activity is responsible for the presence of Sb (III) 2 1. In the same study, reducing conditions, including the presence of H 2 S , did not result in significant reduction of Sb (V) to Sb (III) in water, although the formation of Sb(III)-S compounds was postulated 2 1. In the present study, water from Meager Creek was sampled from locations near microbial mats, which have been shown to exist under reducing conditions (Chapter 4) and to produce MesSb 2 2 . Reducing conditions and microbial metabolism may lead to the presence of Sb (III) in these waters. 6.3.1.2. Methylated antimony species Methylated antimony species were detected in a few samples by using H G - G C - A A S . Biota samples containing methylantimony species include Drepanocladus sp. from Y K locations 1 and 4 and snails (Stagnicola sp.) from Y K locations 1+3 (Table 6.3a). A chromatogram showing stibine, dimethylstibine and trimethylstibine that were generated from a standing water sample ( Y K location 4) is shown in Figure 6.2. 247 V o I t s 0 .14 0.12 o . i q o . o a 0.06\ 0.04 0.02 o . o q - 0 . 0 2 C . 1 4 Me2SbH 1.9 min 0 0.5 1.0 1.5 2.0 2.5 time (minutes) 3.0 3.5 4.0 Figure 6.2. Chromatogram obtained by using HG-GC-AAS (217.6 nm) showing stibines generated at neutral pH from 100 mL of a sample of standing water from location 4 in Yellowknife. 248 The moss samples from Yellowknife that contained methylantimony compounds were all identified as Drepanocladus sp. Interestingly, no detectable antimony species could be extracted from another sample of Drepanocladus sp., at Y K location 15 (see Figure 6. lb). The inability to extract antimony in Drepanocladus sp. from Y K location 15 is probably because of lower levels of total antimony in this sample. The discovery of methylantimony compounds in snails represents, to our knowledge, the first finding of methylated antimony in an animal. Methylantimony compounds in biota have been observed rarely in past studies. Dodd et al. identified predominantly a trimethylantimony compound in one sample and a dimethylantimony compound in another sample of the same species of macrophyte from Yellowknife 5 . The results from the present study differ from that by Dodd et al.5 because the dimethylantimony compound is possibly the same in all three moss samples. Analysis of a soil extract by using H P L C - I C P - M S indicated the presence of a MesSb(V) species7. A dimethylantimony compound is present in a water sample from Y K location 4 and a trimethylantimony compound is present in a water sample from M C location S2 (Table 6.3b). Monomethylantimony and dimethylantimony compounds, assumed to be methylstibonic acid and dimethylstibinic acid, have been observed in river, estuary and sea water samples by other researchers and their presence was attributed to biological activity 2 1 ' 2 3 . The dimethylantimony compound present in the Yellowknife water sample ( Y K location 4) may be similar to the dimethyl compound in the moss sampled from the same location ( Y K location 4), and its presence in both samples may indicate uptake and/or excretion. From these results it is impossible to differentiate between the possibility of the moss forming a dimethylantimony species as a metabolite and excreting it into the water, and the possibility of microorganisms in the water or sediment forming the compound and its being taken up by the moss. Both scenarios are possible. 249 The presence of trimethylantimony species in Meager Creek water may be a result of the production of M e 3 S b by the microbial mats nearby. Recent studies have indicated that Me 3 Sb would be oxidized rapidly to M e 3 S b 0 2 4 ' 2 5 , but other studies suggest that i f the Challenger mechanism is followed for the methylation of antimony (oxidative addition of methyl groups, followed by reduction to the stibine, see Chapter 1, Figure 1.1) the final reduction step to Me 3 Sb takes place only to a very small extent9. Therefore the trimethylantimony compound in the Meager Creek water sample may be present as a result of oxidation following M e 3 S b production in the mats, or of biological antimony methylation only as far as Me 3 SbO. 6.3.1.3. Extraction efficiencies for biota and percent Sb species of total Sb in waters Extraction efficiencies of antimony from biota range from 0.7 to 95%. However, extraction efficiencies for all samples except one are below 37% (Table 6.3a). Clearly the extraction method using MeOH/water (1:1) inadequately extracts antimony from these samples. The reasons for this are likely similar to those given for low extraction efficiencies of arsenic from biota: antimony may be strongly bound to cellular components such as lipids, cellulose, lignin or carbohydrates. For example, metals are known to bind strongly to fungal cell walls 2 6 , and antimony may thus be strongly bound to cellular components. Accordingly, extraction efficiencies from fungus samples (lichen and mushrooms) ranged from 0.8 to 15% (Table 6.3a). Typhus sp. (cattail) from Y K location 12 was extracted nearly quantitatively (Table 6.3a). Some of the antimony extracted from these samples may not have been detected by using H G - G C - A A S , which may account for the differences in amounts detected in waters by H G - G C -A A S compared with the levels of total antimony in waters (Table 6.3b, last column). Arsenic species that are "hidden" to H G - G C - A A S detection, without strong digestion techniques (e.g., U V photolysis or microwave digestion), have been found in sediment pore water samples from 250 Yellowknife . This arsenic was suggested to be bound to colloidal organic matter, or in the form of organoarsenic compounds (such as arsenocholine or arsenosugars)27. In the same way, antimony that is complexed strongly to organic groups may not form hydrides under the conditions used, or Sb-S compounds may exist as well. Sb(III)-S compounds have been proposed to be present in estuarine and interstitial waters, detected by acidifying and degassing samples before H G - G C - A A S analysis at p H 6 2 1 . This method was not carried out in the present study and hence the possibility has not been ruled out that such compounds are present. 6.3.1.4. Total concentrations of antimony compared with arsenic The total concentrations of arsenic (from Chapters 4 and 5) and antimony (from Table 6.3) in some Yellowknife and Meager Creek biota and water samples are summarized in Table 6.4. A n average [Sb]/[As] ratio of 6.4% can be calculated for the biota samples. For water samples other than those taken from Baker Creek (locations 1 and 3) and the effluent from Con Mine (location 13), the average ratio is 4.0%. 251 Table 6.4. Comparison of total concentrations of antimony and arsenic (ppm unless otherwise stated) for selected samples Sample [Sb] [As] [Sb]/[As] x 100% Y K Biota samples (location #) Moss 1, June (5) 190 1130 17 Drepanocladus sp., June (1) 12 1220 9.8 Drepanocladus sp., August (1) 60 490 12 Drepanocladus sp., August (4) 28 880 3.2 Stagnicola sp., August (1+3) 6 82 7.3 Typha sp. (12) 0.20 3.8 5.3 Bidens cernua (10) 0.7 100 0.7 Lemna minor (10) 0.39 28 1.4 Myriophyllum sp., August (10) 0.28 17.4 1.6 Potomogetan richardsonii (6) 0.8 20 4.0 Sparganium augustifolium (6) 0.26 2.5 10 Lichen 4 (7) 120 2300 5.2 Cladonia sp. (13) 1.4 29 4.8 Lycoperdon sp. (7) 60 1010 5.9 Coprinus sp. (8) 34 410 8.3 M C Biota samples (location #) Mimulus sp. (1+2) 0.5 8.7 5.7 Y K Water Samples in ppb Location 10 3.7 68 5.4 Location 11 1.8 53 3.4 Location 4 50 740 6.8 Location 1 260 120 220 Location 3 380 220 170 Con effluent, location 14 31 51 61 M C Water Samples in ppb Location 2, 5 277 1.8 Location S2 5 237 2.1 Location 4 12 288 4.2 252 Ratios of [Sb]/[As] in soil and rocks average about 1 0 % M and therefore it may appear that uptake of antimony by biota is slightly less than that of arsenic, a phenomenon that has been observed before4. A slight deviation of the average ratio obtained (6.4%) from the typical crustal ratio (10%) might also be observed i f soil ratios were lower than 10% in the Yellowknife environment. The ratio in waters is slightly lower (4%) than the ratio in biota. Although these slight differences are noted, the results are still within an order of magnitude of the typical relative levels of antimony and arsenic in the natural environment. It is interesting to note that the concentrations of antimony in waters from Baker Creek ( Y K locations 1,2 and 3) and in Con Mine effluent ( Y K location 14) are similar to those of arsenic. The probable reason for this is that arsenic is effectively removed by alkaline precipitation with ferric sulfate from the effluents, before the effluents are discharged into the environment2 8, whereas antimony is not. Co-precipitation of arsenic with ferric hydroxides is postulated to take place during effluent treatment, and it has been observed that antimony is removed 10 times less efficiently than arsenic from solution by co-precipitation with ferric and manganese hydroxides 1 7. 6.3.2. The confirmation of antimony in samples containing methylantimony compounds by using HG-GC-AAS The presence of methylantimony species in samples was confirmed by two methods: (a) by using H G - G C - A A S at a different wavelength to corroborate that peaks are due to antimony compounds and not the result of spectral interferences, and (b) by using H G - G C - M S to confirm the structure and presence of hydrides derived from samples (next section, 6.3.3). The three moss samples, from Y K locations 1 and 4, as well as the water sample from Y K location 4, were analyzed by using H G - G C - A A S with the A A S operating at a wavelength of 231.4 nm, which is a 253 secondary absorption line specific to antimony. Peaks appeared at the same retention times as those found at a wavelength of 217.6 nm, and in similar abundances, as summarized in Table 6.5. The snail extract was not analyzed because of limited sample size. Table 6.5. Relative amounts (% of sum of methyl species, estimated by normalizing area counts) for methyantimony peaks in moss and water samples. "Sample ( Y K location^)"™"™"™ Sb (III) (RT=0.7) Me 2 Sb- (RT=1.9) Me 3 Sb-(RT=2.3) A A S at 231.4 nm Drepanocladus sp. June ( 1 ) 1 2 88 0 Drepanocladus sp., Aug. (1) 10 90 0 Drepanocladus sp., Aug. (4) 5 95 0 Water, Aug. (4) 17.5 59 23.5 A A S at 217.6 nm Drepanocladus sp., June ( 1 ) 1 7 83 0 Drepanocladus sp., Aug. (1) 23 77 0 Drepanocladus sp., Aug. (4) 6 94 0 Water, Aug. (4) 17.5 59 23.5 6.3.3. The use of headspace HG-GC-MS for the speciation of antimony compounds MesSb was generated in sealed vials from standard solutions of MesSbCb. by using hydride generation methodology. A sample of the headspace was injected into the G C - M S . A detection limit of 0.08 ng Sb for Me 3 Sb was obtained, corresponding to 1 ng Sb in solution before derivatization. However, the analysis of headspace following hydride generation suffers from imprecision, since R S D values no better than 20% were observed for 5 replicate analyses. 254 4e+4 3e+4 3e+4 2e+4 •g 2e+4 H 8 1e+4 1e+4 H 5e+3 Oe+0 H 6.3a 15 Me 3 Sb 30 1 45 time (s) I 60 75 90 o o 5e+3 4e+3 3e+3 2e+3 1e+3 -A 6.3b S b + 121 123 151 136 M e S b + 138 Oe+0 l l l l l i M i i i i l l l l l l i i , , ! M e 2 S b+ 153 ^ [Me 3 Sb-H]+ 167 110 120 130 140 150 160 170 180 190 m/z Figure 6.3a. Total ion chromatogram resulting from headspace-HG-GC-MS analysis showing Me 3 Sb generated from 30 ng Me3SbCl2 (neutral). Figure 6.3b. Mass spectrum at 54.60 s corresponding to M e 3 S b . 255 In Figure 6.3, a chromatogram and mass spectrum are shown, corresponding to standard trimethylstibine (30 ng Sb as Me 3 Sb) generated by using 30 p L of 1000 ppb M e 3 S b C l 2 solution with 1 m L of deionized water and 0.5 mL of 6% (w/v) N a B F L solution. The dominant characteristic of all mass spectra involving antimony compounds is the appearance of the isotopic pattern due to masses of 121 and 123 (naturally occurring at about 52:48) in all Sb-containing fragments. This isotopic pattern is observed in Figure 6.3 at m/z 165/167, corresponding to [ M e 3 m S b - H ] 7 [ M e 3 1 2 3 S b - H ] + ; at m/z 151/153 corresponding to M e 2 1 2 1 S b + / M e 2 1 2 3 S b + ; at m/z 136/138, corresponding to M e 1 2 1 S b + / M e 1 2 3 S b + ; and at m/z 121/123 corresponding to 1 2 1 S b 7 1 2 3 S b + . This mass spectrum is similar to one obtained previously by using the same G C -M S 2 9 and also to spectra obtained by using a quadrupole mass spectrometer5. The loss of a methyl group from methylstibines is the prevalent fragmentation pattern and is considered to be typical for methylated organometallic compounds 3 0. A s discussed in Chapter 2, enhanced demethylation of trimethylstibine is observed when the hydride generation reaction is performed at low pH. Therefore, acidic H G conditions resulting in demethylation can be used to generate mass spectra for methylstibine and dimethylstibine. In Figure 6.4, the chromatogram obtained from hydride generation of M e 3 S b C l 2 (100 ng Sb) at low p H (Figure 6.4a) and the mass spectra obtained for the peaks assumed to be methylstibine (Figure 6.4b) and dimethylstibine (Figure 6.4c) are shown. The fragmentation pattern for dimethylstibine is similar to that observed for trimethylstibine, because the most abundant m/z appears as a result of methyl loss (MeSb + from Me 2 SbH) . The mass spectrum for the peak at a retention time of 39.27s most likely corresponds to the one expected for M e S b H 2 , even though the high background levels obscures a clear, characteristic fragmentation pattern. However, fragments at m/z 121/123 and 136/138 were observed, corresponding to Sb + and MeSb + , respectively. 256 5e+4 4e+4 3e+4 v> § 2e+4 o o 1e+4 6.4a M e S b h L M e 9 S b H A A J T 15 30 T 45 time (s) 60 75 7e+2 - i 6e+2 -5e+2 -w t— 4e+2 -O o 3e+2 -2e+2 -1e+2 -Oe+0 -90 110 120 130 140 150 160 170 180 190 190 Figure 6.4a. Total ion chromatogram resulting from headspace-HG-GC-MS analysis showing stibines generated from 100 ng M e 3 S b C l 2 (1 M HC1). Figure 6.4b. Mass spectrum at 39.27 s corresponding to M e S b H 2 . Figure 6.4c. Mass spectrum at 49.61 s corresponding to M e 2 S b H . 257 The use of H G - G C - M S was useful in the past for the identification and confirmation of methylated antimony species in plant samples from Yellowknife 5 . The direct injection of fractions of gas samples into an ion trap G C - M S resulted in the conclusive identification of trimethylstibine, trimethylbismuthine and methyltin compounds in landfill and fermentation gases2 9. The headspace H G - G C - M S method developed in this work was anticipated to provide information about the presence of methylstibines following H G of Yellowknife samples. Chromatograms and mass spectra for peaks corresponding in retention time to dimethylstibine for moss samples from Yellowknife are shown in Figures 6.5, 6.6 and 6.7. The mass spectra indicate that the compound found in the headspace following hydride generation of the sample extracts is indeed dimethylstibine, by comparison with the mass spectrum shown in Figure 6.4c. The chromatogram and mass spectra for the peaks corresponding in retention time to dimethylstibine and trimethylstibine for the snail extract are shown in Figure 6.8. Again, comparison of the mass spectra with those for standards (Figures 6.3b and 6.4c) indicates the presence of dimethyl- and trimethylstibine following H G of the extract. Differences in m/z abundances are probably due to interfering ions causing slightly different fragmentation patterns. For example, in Figure 6.8c (the mass spectrum for the peak corresponding to trimethylstibine) m/z 166 and 168 are observed (corresponding to Me3Sb+) rather than 165 and 167 (corresponding to [Me3Sb-H]+), which were the m/z observed for the standard compound (Figure 6.3b). For this sample, background subtraction was necessary to isolate the major m/z of interest, because of low levels of antimony in the extract and large amounts of other matrix components. 258 2.5e+4 -j 6.5a 2.0e+4 -1.5e+4 - ]| Me2SbH I | 1 .Oe+4 - \ A o \ l \ 5.0e+3 -O.Oe+0 -i i i i i 0 15 30 45 time (s) 60 75 90 1.6e+3 1.4e+3 -1.2e+3 -1 .Oe+3 CO § 8.0e+2 H o o 6.0e+2 -4.0e+2 -2.0e+2 -0.0e+0 6.5b 110 Sb+ 121 121 136 MeSb+ 138 lilili Me2Sb+ ^153 1111,1111 I I I I | M . . I H I I 120 130 140 150 m/z 160 170 180 190 Figure 6.5a. Total ion chromatogram resulting from headspace-HG-GC-MS analysis of 5 mL of moss extract (June, YK Location 1) showing Me2SbH. Figure 6.5b. Mass spectrum at 46.90 s corresponding to Me2SbH. 259 1.60+4 - i 1.4e+4 -1.2e+4 -LOe+4 -if) r~ 8.0e+3 -u_ O O 6.0e+3 -4.0e+3 -2.0e+3 -0.0e+0 -45 time (s) 1.4e+3 1.2e+3 -1 .Oe+3 -« 8.0e+2 -c .3 ° 6.0e+2 -4.0e+2 -2.0e+2 0.0e+0 6.6b Sb + i l l 122 136 MeSb+ is ! Me2Sb+ IM11111•111111111.... 111 110 120 130 140 150 160 170 180 190 m/z Figure 6.6a. Total ion chromatogram resulting from headspace-HG-GC-MS analysis of 5 mL of moss extract (August, YK Location 1) showing Me2SbH. Figure 6.6b. Mass spectrum at 46.96 s corresponding to Me2SbH. 260 to c o o 2e+4 1e+4 H 1e+4 H 5e+3 Oe+0 H Figure 6.7a. Total ion chromatogram resulting from headspace-HG-GC-MS analysis of 5 m L of moss extract (August, Y K Location 4) showing Me2SbH. F igure 6.7b. Mass spectrum at 46.91 s corresponding to M e 2 S b H . 261 1e+3 H Oe+0 H 6.8a Me.SbH I Me3Sb IVj 3e+2 H | 2e+2 H o o 1e+2 Oe+0 1 15 30 45 time (s) 60 1 3 6 ^ 6.8b Sb+ MeSb+ ^ 1 3 8 i i u 151 Me2Sb+ 75 153 l-lll ,'• I ••' i i . l n l l l 9to 110 120 130 140 150 160 170 180 190 3e+2 2e+2 H 1e+2 H Oe+0 6.8c 150. Sb+ 121 MeSb+ 136 n i l ^ 1 3 8 I.I I. I [Me2Sb-H]+ 152 Me3Sb+ L U i 166 \ 168 i i 1 1 r 110 120 130 140 150 160 170 180 190 m/z Figure 6.8a. Chromatogram (sum of Sb containing ions) resulting from headspace-HG-GC-MS analysis of 10 mL of snail extract (YK Location 1+3) showing Me2SbH and Me3Sb. Figure 6.8b. Background corrected mass spectrum at 46.96 s corresponding to Me2SbH. Figure 6.8c. Background corrected mass spectrum at 54.05 s corresponding to Me3Sb. 262 6.4. Summary Antimony was extracted from environmental biota samples from Yellowknife and Meager Creek, with extraction efficiencies ranging from 0.7 to 37% for all samples except for Typha sp., from which 95% of antimony was extracted. Total antimony levels in most samples, when compared with arsenic levels, reflected the relative abundance of naturally occurring antimony (about 5-10% of arsenic). The exceptions were water samples from Baker Creek, receiving mine effluent from Giant Mine in Yellowknife, which contained antimony at concentrations similar to those for arsenic. This probably indicates that treatment of effluent to remove arsenic is successful, but that antimony is inefficiently removed during the treatment process. Speciation analysis was carried out by using H G - G C - A A S . The major antimony species in all samples, including biota extracts and water, was Sb (V). Sb (III) and methylated antimony species were detected in some samples as well. The presence of methylated antimony species in moss from Yellowknife and a water sample from Yellowknife was confirmed by using H G - G C -A A S at a second absorption wavelength, increasing the likelihood that the peaks obtained are due to the presence of antimony compounds. A headspace H G - G C - M S method was developed for the speciation of antimony compounds and this was used to confirm methylantimony species in the headspace following H G of extracts of moss and snail samples from Yellowknife. Because of the abundance of Drepanocladus sp. at location 1 in Yellowknife, and the seasonal consistency in its methylantimony content, this species of moss can be used as a laboratory standard for dimethylantimony. Future work could involve the use of moss extracts to study H P L C behaviour of the dimethylantimony compound present in the moss by using I C P - M S detection or M S detection for structural information. References 1. Maeda, S. In The chemistry of organic arsenic, antimony and bismuth compounds, Patai, S. Ed. ; John Wiley & Sons: Chichester, 1994; pp 737-742. 2. Keith, L . H . ; Telliard, W . A . Environ. Sci. Technol. 1919,13, 416. 3. Cutter, G . A . ; Cutter, L . S. Mar. Chem. 1995, 49, 295-306. 4. Stewart, K . C ; M c K o w n , D . M . J. Geochem. Explor. 1995, 54, 19-26. 5. Dodd, M . ; Pergantis, S. A . ; Cullen W . R.; L i , H . ; Eigendorf, G . K . ; Reimer, K . J. Analyst 1996, 121, 223-228. 6. Lintschinger, J.; Koch , I.; Serves, S.; Feldmann, J.; Cullen, W . R. Fresenius J. Anal. Chem. 1997, 359, 484-491. 7. Ulrich, N . Anal. Chim. Acta 1998, 359, 245-253. 8. Smichowski, P.; Madrid, Y . ; De L a Calle Guntinas, M . B . ; Camara, C. J. Anal. At. Spectrom. 1995, 10, 815-821. 9. Andrewes, P.; Cullen, W. R.; Feldmann, J.; Koch, I.; Polishchuk, E . Appl. Organomet. Chem. 1998, in press. 10. Koch, I.; Feldmann, J.; Lintschinger, J.; Serves, S. V . ; Cullen, W . R.; Reimer, K . J. Appl. Organomet. Chem. 1998, 12, 129-136. 11. Dodd, M . ; Grundy, S. L . ; Reimer, K . J.; Cullen, W . R. Appl. Organomet. Chem. 1992, 6, 207. 12. Morgan, G . T.; Davies, G . R. Proc. Royal Soc, Ser. A 1926, 523. 13. Bajo, S.; Suter, U . ; Aeschliman, B . Analytica Chimica Acta 1983, 149, 321-355. 14. Cullen, W. R.; L i , H . ; Hewitt, G . ; Reimer, K . J.; Zalunardo, N . Appl. Organomet. Chem., 1994, 8, 303 15. Le , X . C ; Cullen, W. R.; Reimer, K . J Appl. Organomet. Chem. 1992, 6, 161. 16. Kantin R. Limnol. Oceanogr. 1983, 28, 165-168. 17. M o k , W . - M . ; Wai, C. M . Environ. Sci. Technol. 1990, 24, 102-108. 264 18. Mohommad, B . ; Ure, A . M . ; Reglinski, J.; Littlejohn, D . Chem. Speciation Bioavail. 1990, 3, 117-122. 19. Yamamoto, M . ; Urata, K . ; Murashige, K . ; Yamamoto, Y . Spectrochim. Acta 1981, 36B, 671-677. 20. Pourbaix, M . Atlas of Electrochemical Equilibria in Aqueous Solutions; National Association of Corrosion Engineers: Houston, Texas, 1974; p 524-532. 21. Bertine, K . K . ; Lee, D . S. In Trace Metals in Seawater, NATO Conference Series, Series IV: Marine Sciences; Wong, C. S.; Boyle, E . ; Bruland, K . W. ; Berton, J. D . ; Goldberg, E . D. , Eds.; Plenum: N e w York, 1983; pp 21-38. 22. Feldmann, J.; Lehr, C ; Koch, I; Andrewes, P.; La i , V . W . - M . ; Cullen, W . R., manuscript in preparation. 23. Andreae, M . O.; Asmode, J.-F.; Foster, P.; Van't dack, L . Anal. Chem. 1981, 53, 1766-1771. 24. Parris, G. E . ; Brinckmann, F. E . Environ. Sci Tech. 1976, 10, 1128. 25. Jenkins, R. O.; Craig, P. J.; Goessler, W. ; Miller, D . Ostah, N . ; Irgolic, K . J. Environ. Sci. Tech. 1998, 32, 882-885. 26. Sarret, G . ; Manceau, A . ; Spandini, L . ; Roux, J . - C ; Hazemann, J . -L. ; Soldo, Y . ; Eybert-Berard, L . ; Menthonnex, J.-J. Environ. Sci. Technol. 1998, 32, 1648-1655. 27. Bright, D . A . ; Dodd, M . ; Reimer, K . J. Sci. Tot. Environ. 1996, 180, 165-182. 28. Halverson, G . B . ; Raponi, R. R. Water Poll. Res. J. Canada. 1987, 22, 570-583. 29. Feldmann, J.; Koch , I.; Cullen, W. R. Analyst, 1998, 815-820. 30. Nekrasov, Y . S.; Zagorevskii, D . V . In The Chemistry of Organic Arsenic, Antimony and Bismuth Compounds; Patai, S., Ed . ; John Wiley: Chichester, 1994; pp 237-264. 265 Chapter 7 CONCLUSIONS AND FUTURE WORK Considerable knowledge was gained about arsenic and antimony species in the terrestrial environment. Existing methods for speciation analysis were adapted for this work. Three H P L C methods with I C P - M S detection allowed separation and identification of at least 11 arsenic species. The agreement of retention times between chromatographic peaks in samples and standards, by using more than one method, corroborates identifications based on retention times. Arsenosugar standards were analyzed by using tandem ESI - IT-MS and fragmentation patterns specific to this mass analyzer were obtained. Only partial identification of arsenosugars in a crude kelp extract was achieved by using M S - M S and M S - M S - M S techniques (only arsenosugar XIII could be identified with any certainty). Future work should address clean-up of the kelp extract (e.g., the use of H P L C methods, before introduction to ESI- IT-MS) . The demethylation of trimethylantimony species during analysis by using H G - G C -A A S was characterized as being dependent on pH; lower p H resulted in more demethylation. Mechanistic reasons for demethylation were sought and it was established that methyl groups were lost during the H G reaction, rather than as a result of instability of the starting compound (Me3SbCl2) or the product (Me 3 Sb) to acid. Care is recommended (and was taken) in the analysis of environmental or other samples for 266 antimony by using H G - G C methods to ensure that the methylantimony species identified are not an artifact of the method. Non-toxic and very low levels of toxic water soluble arsenic species were identified in some edible mushrooms, and consequently these do not present a health concern to consumers. It would be interesting to determine i f arsenic speciation changes as the arsenic concentration increases for the mushrooms analyzed. In some cases the extractable arsenic was inorganic and hence higher levels might become of greater toxicological concern. Pure culture experiments with fungi that can produce mushrooms indicated that the synthesis of arsenobetaine, arsenocholine or arsenosugars does not take place by the mycelia during the short experiment times. Arsenobetaine is accumulated by Scleroderma citrinum which may suggest that i f any arsenobetaine is present in the growing environment of wild mushrooms, it may be accumulated by the fungus. The interaction of antimony with fungus was confounded by the ubiquitous presence of an unknown antimony compound for which only partial characterization was accomplished. Pleurotus flabellatus formed an antimony-containing metabolite (of unknown identity, detected by using H P L C - I C P - M S ) from inorganic antimony. This fungus also oxidized Sb (III) to Sb(OH) 6". Future experiments should include growth media that contain a minimum amount of salts and/or carbon sources, to simplify the matrix and hopefully eliminate the presence of unidentified antimony-binding compounds. Novel results were obtained from a study of the arsenic species in samples from two terrestrial environments: a hot springs environment (Meager Creek, B C ) and from an area impacted by smelting and mining (Yellowknife, N W T ) . Arsenosugars are apparently synthesized by cyanobacteria and other bacteria in microbial mats from both locations. 267 Thus thermophilic as well as non-thermophilic organisms have this capability. Small amounts of arsenosugars were also found in lichens, in higher plants, in some mushrooms, in green algae (belonging to the phylum Chlorophyta), in freshwater mussels and in suckers. Because they are bottom feeders, suckers probably accumulate arsenosugars from benthic organisms. Suckers also contain arsenobetaine which was the major arsenic compound found in all other fish analyzed, except for pike. In pike, the major detectable arsenic compound was D M A . On the other hand, neither freshwater mussels nor snails contain appreciable quantities of arsenobetaine. This is in direct contrast to findings in the marine environment where marine mussels and gastropods usually contain arsenobetaine as a major or as the only arsenic compound. Freshwater mussels contain mostly arsenosugars and snails contained mostly tetramethylarsonium ion, as well as inorganic arsenic. Arsenobetaine was found for the first time in lichens and it is present in all specimens from Yellowknife. It was also found in some mushrooms, including Coprinus comatus and Lycoperdon pyriforme. The identity of arsenic species was determined for the first time in Paxillus involutus, Psathyrella candolleana and Leccinum scabrum. The major arsenic species extracted from higher plants, as well as lichens, mosses, algae and microbial mats were inorganic (arsenite and arsenate). A large amount of arsenic remained unextracted or undetected in all types of samples. The nature of this arsenic is unknown and hence more studies are imperative to determine its chemical and toxicological significance. Arsenic that is not extracted may be bound to lipids, cell components, proteins, or it may exist in a mineral form. 2 6 8 Antimony species were determined in some samples from Meager Creek and Yellowknife, and mostly Sb (V) was found (although, as for arsenic, a large fraction remained unextracted). In moss samples, however, a dimethylantimony species in the form of a compound that resulted in dimethylstibine being formed from hydride generation of the extract, is present. Derivatization of extracts of the same moss species from two different locations, and from the same location at different times (June and August) yielded dimethylstibine in all cases. Future work to identify this compound would be very interesting. Arsenic speciation in the terrestrial environment, in general, does not appear to be as complex as in the marine environment. For example, arsenosugars and arsenobetaine, which are the major arsenic compounds in marine plants and animals, respectively, occur either rarely or in very small amounts in terrestrial samples. This may indicate that different metabolic pathways are followed in the terrestrial environment, or that the processes in the marine environment are not important in the terrestrial environment. Little is known about the speciation of antimony in the environment, and this study helps to establish a knowledge base for this element. Inorganic antimony appears to be the predominant species and there is no evidence yet for antimony analogues of arsenosugars or arsenobetaine. However the apparent absence of these compounds may be due to the present lack of appropriate analytical methodology for antimony speciation. 269 

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