Determination of Organic Compounds in Natural and Treated Waters
Determination of Organic Compounds in Natural and Tr...
54 downloads
1729 Views
4MB Size
Report
This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!
Report copyright / DMCA form
Determination of Organic Compounds in Natural and Treated Waters
Determination of Organic Compounds in Natural and Treated Waters
T.R. Crompton
London and New York
First published 2000 by E & FN Spon 11 New Fetter Lane, London EC4P 4EE Simultaneously published in the USA and Canada by E & FN Spon 29 West 35th Street, New York, NY 10001 E & FN Spon is an imprint of the Taylor & Francis Group This edition published in the Taylor & Francis e-Library, 2002. © 2000 T.R. Crompton All rights reserved. No part of this book may be reprinted or reproduced or utilized in any form or by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying and recording, or in any information storage or retrieval system, without permission in writing from the publishers. The publisher makes no representation, express or implied, with regard to the accuracy of the information contained in this book and cannot accept any legal responsibility or liability for any errors or omissions that may be made. British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library Library of Congress Cataloging in Publication Data Crompton, T.R. (Thomas Roy) Determination of organic compounds in natural and treated waters/T.R. Crompton. p. cm. Includes bibliographical references and index. 1. Water—Analysis. 2. Organic compounds. I. Title. QD142.C76 1999 628.1´61—dc21 98–52383 CIP ISBN 0 419 24360 7 (Print Edition) ISBN 0-203-01635-1 Master e-book ISBN ISBN 0-203-13987-9 (Glassbook Format)
Contents
Preface 1
Introduction
1.1 Brief summary of methodologies 1.1.1 Titration procedures 1.1.2 Polarographic methods 1.1.3 Spectrophotometric methods 1.1.3.1 Visible spectrometry 1.1.3.2 Ultraviolet spectrometry 1.1.3.3 Luminescence and fluorescence spectrometry 1.1.3.4 Infrared and Raman spectrometry 1.1.4 Flow injection analysis 1.1.5 Atomic spectrometric methods 1.1.5.1 Atomic absorption spectrometry 1.1.5.2 Inductively coupled plasma atomic emission spectrometry 1.1.5.3 Atomic emission spectrometry 1.1.6 Nuclear magnetic resonance spectrometry 1.1.7 Neutron activation analysis 1.1.8 X-ray fluorescence spectrometry 1.1.8.1 Energy-dispersive X-ray fluorescence spectrometry 1.1.8.2 Total reflection X-ray fluorescence spectrometry 1.1.9 Isotope dilution methods 1.1.10 Enzymic assay methods 1.1.11 Chromatographic methods 1.1.11.1 Gas chromatography 1.1.11.2 Gas chromatography–mass spectrometry 1.1.11.3 Headspace gas chromatography
xlii 1 1 1 2 5 5 7 7 12 13 15 15 17 24 24 24 25 25 27 33 33 33 34 43 52
vi
Contents
1.1.11.4 1.1.11.5 1.1.11.6 1.1.11.7 1.1.11.8 1.1.11.9 1.1.11.10 1.1.11.11 1.1.11.12 1.1.11.13
Purge and trap gas chromatography Pyrolysis–gas chromatography High-performance liquid chromatography High-performance liquid chromatography– mass spectrometry Ion-exchange chromatography Column coupling capillary isotachoelectrophoresis Ion chromatography Conventional column chromatography Supercritical fluid chromatography Thin-layer chromatography
1.2 Preconcentration techniques 1.2.1 Introduction 1.2.2 Review of preconcentration techniques 1.2.3 Analytical finishes after preconcentration
54 54 56 65 66 67 67 70 71 75 77 77 81 81
1.3 Rationale, analysis of water samples References
81 105
2
109
Hydrocarbons
2.1 Aliphatic hydrocarbons 2.1.1 Natural (non-saline) waters 2.1.1.1 Gas chromatography 2.1.1.2 Headspace analysis 2.1.1.3 Gas stripping methods 2.1.1.4 High-performance liquid chromatography 2.1.1.5 Infrared spectroscopy 2.1.1.6 Thin-layer chromatography 2.1.1.7 Paper chromatography 2.1.1.8 Miscellaneous 2.1.2 Sea and estuary waters 2.1.2.1Gas chromatography 2.1.2.2Headspace analysis 2.1.2.3Gas stripping analysis 2.1.2.4Miscellaneous 2.1.3 Potable waters 2.1.3.1Gas chromatography 2.1.3.2Preconcentration
110 110 110 111 112 115 116 123 125 125 126 126 126 127 127 128 128 129
Contents
2.1.4
vii
Waste waters 2.1.4.1 Infrared spectrometry 2.1.4.2 Thin-layer chromatography 2.1.4.3 Miscellaneous Trade effluents 2.1.5.1 Spectrophotometric methods 2.1.5.2 Miscellaneous 2.1.5.3 Preconcentration Sewage effluents 2.1.6.1 Gas chromatography 2.1.6.2 Miscellaneous Petroleum and diesel fuel spills 2.1.7.1 Gas chromatography 2.1.7.2 Gas stripping methods 2.1.7.3 Thin-layer chromatography 2.1.7.4 Column chromatography 2.1.7.5 Infrared spectrometry 2.1.7.6 Ultraviolet spectrometry 2.1.7.7 Miscellaneous
129 129 132 134 135 135 135 136 136 136 136 137 137 137 137 138 139 139 140
2.2 Aromatic hydrocarbons 2.2.1 Natural waters 2.2.1.1 Spectrophotometric method 2.2.1.2 Gas chromatography 2.2.1.3 Infrared spectrometry 2.2.1.4 Column chromatography 2.2.1.5 Thin-layer chromatography 2.2.1.6 Ultraviolet spectroscopy 2.2.1.7 Miscellaneous 2.2.1.8 Preconcentration 2.2.2 Trade effluents 2.2.2.1 Gas chromatography
140 140 141 141 142 142 142 142 144 144 144 144
2.3 Polyaromatic hydrocarbons 2.3.1 Natural (non-saline) waters 2.3.1.1 Gas chromatography 2.3.1.2 High-performance liquid chromatography 2.3.1.3 Thin-layer chromatography 2.3.1.4 Fluorescence spectrometry 2.3.1.5 Extraction of polyaromatic hydrocarbons from water
145 145 146 152 155 155
2.1.5
2.1.6
2.1.7
157
viii
Contents
2.3.2 2.3.3 2.3.4
2.3.5
2.3.6
2.3.7
2.3.1.6 Miscellaneous 2.3.1.7 Preconcentration Aqueous precipitation 2.3.2.1 Miscellaneous Sea water 2.3.3.1 Fluorescence spectrometry Potable water 2.3.4.1 Gas chromatography 2.3.4.2 Thin-layer chromatography 2.3.4.3 High-performance liquid chromatography 2.3.4.4 Fluorescence spectrometry 2.3.4.5 Polarography 2.3.4.6 Miscellaneous 2.3.4.7 Preconcentration Waste waters 2.3.5.1 Gas chromatography 2.3.5.2 High-performance liquid chromatography 2.3.5.3 Low-temperature spectrofluorimetry using the Shpol’skii effect Trade effluents 2.3.6.1 Gas chromatography 2.3.6.2 High-performance liquid chromatography 2.3.6.3 Miscellaneous 2.3.6.4 Preconcentration Sewage effluents 2.3.7.1 High-performance liquid chromatography 2.3.7.2 Miscellaneous
2.4 Mineral oil and petroleum spills 2.4.1 Non-saline natural waters 2.4.1.1 Introduction 2.4.1.2 Gas chromatography 2.4.1.3 Thin-layer chromatography 2.4.1.4 Infrared spectrometry 2.4.1.5 Fluorescence techniques 2.4.1.6 Characterization of oils by metals analysis 2.4.1.7 Miscellaneous 2.4.1.8 Preconcentration 2.4.2 Saline waters 2.4.2.1 Gas chromatography 2.4.2.2 Paper chromatography
157 157 158 158 158 158 158 159 161 163 166 170 170 170 174 174 174 175 175 175 177 177 177 178 178 178 178 178 178 183 189 193 194 197 199 201 201 201 210
Contents
2.4.3 2.4.4 2.4.5
2.4.2.3 Gel permeation chromatography 2.4.2.4 Infrared spectroscopy 2.4.2.5 Raman spectroscopy 2.4.2.6 Ultraviolet spectroscopy 2.4.2.7 Fluorescence spectrometry 2.4.2.8 Characterization of oils by metals analysis 2.4.2.9 Miscellaneous Waste waters 2.4.3.1 Gas chromatography Trade effluents 2.4.4.1 Fluorescence spectrometry Sewage effluents 2.4.5.1 Miscellaneous
ix
210 211 214 214 214 216 216 217 217 217 217 218 218
2.5 Mineral greases
218
2.6 Vegetable oils References
219 219
3
232
Oxygen-containing compounds
3.1 Carboxylic acids 3.1.1 Natural (non-saline) waters 3.1.1.1 Spectrophotometric methods 3.1.1.2 Gas chromatography 3.1.1.3 Thin-layer chromatography 3.1.1.4 High-performance liquid chromatography 3.1.1.5 Ion-exclusion chromatography 3.1.1.6 Ion chromatography 3.1.1.7 Fluorescence spectrometry 3.1.1.8 Potentiometric methods 3.1.1.9 Differential pulse polarography 3.1.1.10 Miscellaneous 3.1.1.11 Preconcentration 3.1.2 Rain water 3.1.2.1 Gas chromatography 3.1.2.2 High-performance liquid chromatography 3.1.2.3 Ion-exclusion chromatography 3.1.3 Sea water 3.1.3.1 Gas chromatography 3.1.3.2 Atomic absorption spectrometry 3.1.3.3 High-performance liquid chromatography
232 232 232 234 235 236 238 238 240 240 240 240 241 241 241 241 241 242 242 242 242
x
Contents
3.1.3.4 Diffusion method Waste waters 3.1.4.1 Gas chromatography 3.1.4.2 Miscellaneous Trade effluents 3.1.5.1 Spectrophotometric methods 3.1.5.2 Gas chromatography 3.1.5.3 Miscellaneous Sewage effluents 3.1.6.1 Gas chromatography 3.1.6.2 Column chromatography 3.1.6.3 Miscellaneous
243 243 243 243 243 243 244 244 245 245 248 248
3.2 Alcohols 3.2.1 Natural waters 3.2.1.1 Spectrophotometric methods 3.2.1.2 Gas chromatography 3.2.1.3 Miscellaneous
248 248 248 249 249
3.2.2
249 249 249 249
3.1.4
3.1.5
3.1.6
3.2.3
Trade effluents 3.2.2.1 Spectrophotometric method Sewage effluents 3.2.3.1 Gas chromatography
3.3 Glycols 3.3.1 Natural waters 3.3.1.1 Spectrophotometry 3.3.1.2 Gas chromatography
250 250 250 250
3.4 Aldehydes 3.4.1 Natural waters 3.4.1.1 Spectrophotometric methods 3.4.1.2 Gas chromatography 3.4.1.3 Thin-layer chromatography 3.4.1.4 Polarography 3.4.1.5 Preconcentration 3.4.2 Potable water 3.4.2.1 Purge and trap gas chromatography 3.4.3 Trade effluents 3.4.3.1 Gas chromatography 3.4.4 Sewage effluents 3.4.4.1 Spectrophotometric method
250 250 250 251 251 251 251 252 252 252 252 252 252
Contents
xi
3.5 Ketones 3.5.1 Natural waters 3.5.1.1 Gas chromatography
253 253 253
3.6 Esters 3.6.1 Natural waters 3.6.1.1 Gas chromatography 3.6.1.2 High-performance liquid chromatography 3.6.2 Potable waters 3.6.2.1 High-performance liquid chromatography 3.6.3 Waste waters 3.6.3.1 Polarography 3.6.4 Trade effluents 3.6.4.1 Gas chromatography 3.6.4.2 Electrokinetic chromatography 3.6.4.3 Miscellaneous
253 253 253 254 257 257 257 257 257 257 258 258
3.7 Carbohydrates 3.7.1 Natural waters 3.7.1.1 Spectrophotometric method 3.7.1.2 Gas chromatography 3.7.1.3 Enzymic assay 3.7.1.4 Miscellaneous 3.7.2 Sea water 3.7.2.1 Gas chromatography 3.7.2.2 Partition chromatography 3.7.2.3 Miscellaneous 3.7.3 Waste waters 3.7.3.1 Spectrophotometric method 3.7.4 Trade effluents 3.7.4.1 Gas chromatography 3.7.4.2 Gel chromatography
258 258 258 258 258 259 259 259 259 259 261 261 261 261 261
3.8 Lactams 3.8.1 Natural waters 3.8.1.1 Thin-layer chromatography 3.8.1.2 Polarography
261 261 261 262
3.9 Dioxans 3.9.1 Natural waters 3.9.1.1 Spectrophotometric method
262 262 262
xii
Contents
3.10 Quinones 3.10.1 Waste waters 3.10.1.1Thin-layer chromatography 3.10.1.2Polarography 3.10.2 Trade effluents 3.10.2.1 Thin-layer chromatography
263 263 263 263 263 263
3.11 Phenols 3.11.1 Natural waters 3.11.1.1 Spectrophotometric methods 3.11.1.2 Gas chromatography 3.11.1.3 Gas chromatography-mass spectrometry 3.11.1.4 Thin-layer chromatography 3.11.1.5 High-performance liquid chromatography 3.11.1.6 Adsorption of phenols on resin columns 3.11.1.7 Raman spectroscopy 3.11.1.8 Ultraviolet spectroscopy 3.11.1.9 Fluorescence spectrometry 3.11.1.10 Atomic absorption spectroscopy 3.11.1.11 Sample collection and preservation 3.11.1.12 Miscellaneous 3.11.1.13 Preconcentration 3.11.2 Potable waters 3.11.2.1 Spectrophotometric methods 3.11.2.2 Gas chromatography 3.11.2.3 Thin-layer chromatography 3.11.2.4 Raman spectroscopy 3.11.3 Waste waters 3.11.3.1 Spectrophotometric methods 3.11.3.2 Thin-layer chromatography 3.11.3.3 Resin column analysis 3.11.3.4 Miscellaneous 3.11.4 Trade effluents 3.11.4.1 Spectrophotometric methods 3.11.4.2 Gas chromatography 3.11.4.3 High-performance liquid chromatography 3.11.4.4 Ultraviolet spectrometry 3.11.4.5 Miscellaneous 3.11.5 Sewage effluents 3.11.5.1 Spectrophotometric methods 3.11.5.2 Flow injection analysis
263 263 264 268 273 273 274 275 275 277 277 277 278 278 278 279 279 280 280 281 281 281 282 282 282 283 283 283 283 284 284 284 284 285
Contents
3.11.5.3 3.11.5.4 3.11.5.5 3.11.5.6
Gas chromatography High-performance liquid chromatography Enzymic assay Miscellaneous
xiii
285 285 285 286
3.12 Phenolic acids 3.12.1 Natural waters 3.12.1.1 Thin-layer chromatography
288 288 288
3.13 Ethers 3.13.1 Potable waters 3.13.1.1 High-performance liquid chromatography References
289 289 289 289
4
298
Surface active agents
4.1 Anionic detergents 4.1.1 Natural waters 4.1.1.1 Titration methods 4.1.1.2 Spectrophotometric methods 4.1.1.3 Gas chromatography 4.1.1.4 High-performance liquid chromatography 4.1.1.5 Infrared spectrometry 4.1.1.6 Ultraviolet spectroscopy 4.1.1.7 Polarography 4.1.1.8 Potentiometric analysis 4.1.1.9 Atomic absorption spectrometry 4.1.1.10 Miscellaneous 4.1.2 Sea waters 4.1.2.1 Spectrophotometric methods 4.1.2.2 Polarography 4.1.2.3 Atomic absorption spectrometry 4.1.2.4 Miscellaneous 4.1.3 Potable waters 4.1.3.1 Spectrophotometric methods 4.1.3.2 Polarography 4.1.3.3 Miscellaneous 4.1.3.4 Preconcentration 4.1.4 Waste waters 4.1.4.1 Spectrophotometric methods 4.1.4.2 Column chromatography 4.1.4.3 Infrared spectroscopy
298 298 298 299 301 303 304 304 305 305 305 306 307 307 307 307 309 310 310 311 311 312 312 312 312 312
xiv
Contents
4.1.4.4 Preconcentration Sewage effluents 4.1.5.1 Spectrophotometric methods 4.1.5.2 High-performance liquid chromatography 4.1.5.3 Polarography
314 314 314 314 314
4.2 Cationic detergents 4.2.1 Natural waters 4.2.1.1 Titration methods 4.2.1.2 Spectrophotometric methods 4.2.1.3 Flow injection analysis 4.2.1.4 High-performance liquid chromatography 4.2.1.5 Ultraviolet spectroscopy 4.2.1.6 Miscellaneous 4.2.2 Sea water 4.2.2.1 Titration method 4.2.2.2 Atomic absorption spectrometry 4.2.3 Waste waters 4.2.3.1 Flow injection analysis 4.2.4 Biodegradation test liquids 4.2.4.1 Titration method
315 315 315 315 316 316 316 317 317 317 317 317 317 317 317
4.3 Non-ionic detrergents 4.3.1 Natural waters 4.3.1.1 Spectrophotometric methods 4.3.1.2 Gas chromatography 4.3.1.3 Thin-layer chromatography 4.3.1.4 Column chromatography 4.3.1.5 High-performance liquid chromatography 4.3.1.6 Ion-exchange chromatography 4.3.1.7 Polarography 4.3.1.8 Atomic absorption spectrometry 4.3.1.9 Miscellaneous 4.3.1.10 Preconcentration 4.3.2 Sea water 4.3.2.1 Spectrophotometric method 4.3.2.2 Atomic absorption spectrometry 4.3.3 Potable waters 4.3.3.1 Mass spectrometry 4.3.4 Waste waters 4.3.4.1 Spectrophotometric methods
318 318 318 320 322 322 325 325 327 327 329 330 331 331 331 332 332 332 332
4.1.5
Contents
xv
4.3.4.2 Gas chromatography 4.3.4.3 Atomic absorption spectrometry 4.3.4.4 Miscellaneous 4.3.5 Trade effluents 4.3.5.1 Flow injection analysis 4.3.5.2 High-performance liquid chromatography 4.3.6 Sewage effluents 4.3.6.1 Spectrophotometric methods 4.3.6.2 High-performance liquid chromatography References
332 333 333 333 333 334 334 334 334 335
5
339
Halogen-containing compounds
5.1 Saturated aliphatic halogen compounds 5.1.1 Natural waters 5.1.1.1 Gas chromatography 5.1.1.2 Headspace analysis 5.1.1.3 Purge and trap analysis 5.1.1.4 Thin-layer chromatography 5.1.1.5 High-performance liquid chromatography 5.1.1.6 Ion-exchange chromatography 5.1.1.7 Ultraviolet spectrometry 5.1.1.8 Miscellaneous 5.1.1.9 Preconcentration 5.1.2 Sea water 5.1.2.1 Ion-exchange chromatography 5.1.3 Waste waters 5.1.3.1 Gas chromatography 5.1.4 Sewage effluents 5.1.4.1 Gas chromatography 5.1.5 Trade effluents 5.1.5.1 Gas chromatography
339 339 339 342 343 343 344 345 345 345 346 347 347 347 347 348 348 349 349
5.2 Unsaturated aliphatic halogen compounds 5.2.1 Natural waters 5.2.1.1 Gas chromatography 5.2.1.2 Gas chromatography–mass spectrometry 5.2.1.3 Mass spectrometry 5.2.1.4 Headspace analysis 5.2.1.5 Purge and trap analysis 5.2.1.6 High-performance liquid chromatography
349 349 349 350 350 351 351 354
xvi
Contents
5.2.2 5.2.3
5.2.4
Sea water 5.2.2.1 Purge and trap analysis Potable waters 5.2.3.1 Gas chromatography 5.2.3.2 Gas chromatography–mass spectrometry 5.2.3.3 Mass spectrometry 5.2.3.4 Purge and trap method 5.2.3.5 Miscellaneous Trade effluents 5.2.4.1 Gas chromatography
5.3 Haloforms 5.3.1 Natural waters 5.3.1.1 Spectrophotometric method 5.3.1.2 Gas chromatography 5.3.1.3 Headspace analysis 5.3.1.4 Purge and trap methods 5.3.1.5 Resin adsorption–gas chromatography 5.3.1.6 High-performance liquid chromatography 5.3.1.7 Gel permeation chromatography 5.3.1.8 Preconcentration 5.3.2 Sea water 5.3.2.1 Miscellaneous 5.3.3 Potable waters 5.3.3.1 Gas chromatography 5.3.3.2 Gas chromatography–mass spectrometry 5.3.3.3 Static headspace analysis 5.3.3.4 Purge and trap analysis (dynamic headspace analysis) 5.3.3.5 High-performance liquid chromatography 5.3.3.6 Miscellaneous 5.3.3.7 Preconcentration 5.3.4 Trade effluents 5.3.4.1 Solvent extraction–gas chromatography 5.3.4.2 Headspace analysis 5.3.5 Sewage effluents 5.3.5.1 Solvent extraction–gas chromatography 5.3.6 Swimming pool waters 5.3.6.1 Solvent extraction–gas chromatography
354 354 355 355 355 355 355 355 355 355 358 358 358 358 359 362 363 364 364 364 365 365 365 367 383 384 387 392 392 393 393 393 393 393 393 393 393
Contents
xvii
5.4 Haloaromatic compounds 5.4.1 Natural waters 5.4.1.1 Gas chromatography 5.4.1.2 Liquid chromatography 5.4.1.3 Miscellaneous
394 394 394 395 395
5.5 Chlorinated alkyl naphthalenes 5.5.1 Trade effluents 5.5.1.1 Miscellaneous
395 395 395
5.6 Chlorinated polyaromatic compounds 5.6.1 Waste waters 5.6.1.1 High-performance liquid chromatography–gas chromatography
395 395
5.7 Polychlorobiphenyls 5.7.1 Natural waters 5.7.1.1 Gas chromatography 5.7.1.2 Gas chromatography-mass spectrometry 5.7.1.3 Thin-layer chromatography 5.7.1.4 High-performance liquid chromatography 5.7.1.5 Polarography 5.7.1.6 Enzyme immunoassay 5.7.1.7 Miscellaneous 5.7.1.8 Preconcentration 5.7.2 Sea water 5.7.2.1 Gas chromatography 5.7.2.2 High-performance liquid chromatography 5.7.2.3 Miscellaneous 5.7.3 Potable waters 5.7.3.1 Gas chromatography 5.7.4 Trade effluents 5.7.4.1 Gas chromatography 5.7.4.2 Column chromatography 5.7.5 Sewage effluents 5.7.5.1 Gas chromatography 5.7.5.2 Gas chromatography–mass spectrometry 5.7.5.3 Miscellaneous
395 395 396 399 400 401 401 402 402 402 403 403 403 404 404 404 405 405 405 406 406 406 406
5.8 Chlorophenols 5.8.1 Natural waters 5.8.1.1 Spectrophotometric method
407 407 407
395
xviii
5.8.2
5.8.3
5.8.4 5.8.5
Contents
5.8.1.2 Gas chromatography 5.8.1.3 Thin-layer chromatography 5.8.1.4 High-performance liquid chromatography 5.8.1.5 Ion chromatography 5.8.1.6 Polarography 5.8.1.7 Miscellaneous 5.8.1.8 Preconcentration Potable waters 5.8.2.1 Gas chromatography 5.8.2.2 Gas chromatography–mass spectrometry 5.8.2.3 High-performance liquid chromatography Waste waters 5.8.3.1 Gas chromatography 5.8.3.2 High-performance liquid chromatography 5.8.3.3 Preconcentration Trade effluents 5.8.4.1 Gas chromatography–mass spectrometry Sewage effluents 5.8.5.1 Gas chromatography–mass spectrometry 5.8.5.2 Miscellaneous
407 410 410 411 411 411 412 412 412 415 415 415 415 416 417 417 417 417 417 417
5.9 Halocarboxylic acids 5.9.1 Natural waters 5.9.1.1 Gas chromatography 5.9.1.2 Isotope dilution mass spectrometry 5.9.1.3 Miscellaneous 5.9.2 Potable waters 5.9.2.1 Pyrolysis method 5.9.3 Trade effluents 5.9.3.1 Isotachophoresis 5.9.4 Sewage effluents 5.9.4.1 Miscellaneous
417 417 417 418 418 419 419 419 419 419 419
5.10 Chlorinated isocyanurates 5.10.1 Swimming pool water 5.10.1.1 Spectrophotometric method
420 420 420
5.11 Chlorolignosulphonic acids 5.11.1 Natural waters 5.11.1.1Pyrolysis–gas chromatography 5.11.2 Potable waters 5.11.2.1 Pyrolysis–gas chromatography
420 420 420 420 420
Contents
5.11.3
Trade effluents 5.11.3.1 Pyrolysis–gas chromatography
xix
420 420
5.12 Polychlorodibenzo-p-dioxins and polychlorodibenzofurans 5.12.1 Natural waters 5.12.1.1 Gas chromatography–mass spectrometry 5.12.1.2 High-performance liquid chromatography 5.12.2 Potable waters 5.12.2.1 Gas chromatography 5.12.3 Waste waters 5.12.3.1 Miscellaneous
421 421 421 421 422 422 422 422
5.13 Chloroanisoles and chloromethylanisoles 5.13.1 Natural waters 5.13.1.1 Gas chromatography–mass spectrometry
422 422 422
5.14 Polybromobiphenyls 5.14.1 Natural waters 5.14.1.1 Irradiation–gas chromatography
422 422 422
5.15 Polychloroterphenyls 5.15.1 Natural waters 5.15.1.1 Gas chromatography–mass spectrometry References
422 422 422 423
6
434
Nitrogen-containing compounds
6.1 Aliphatic amines 6.1.1 Natural waters 6.1.1.1 Spectrophotometric methods 6.1.1.2 Gas chromatography 6.1.1.3 Gas chromatography–mass spectrometry 6.1.1.4 Thin-layer chromatography 6.1.1.5 Liquid chromatography 6.1.2 Sea water 6.1.2.1 Spectrophotometric methods 6.1.2.2 Miscellaneous 6.1.3 Potable waters 6.1.3.1 Gas chromatography–mass spectrometry 6.1.4 Trade effluents 6.1.4.1 Titration procedure 6.1.4.2 Spectrophotometric method 6.1.4.3 Gas chromatography
434 434 434 434 435 435 435 435 435 435 436 436 436 436 436 436
xx
Contents
6.1.5
6.1.4.4 Thin-layer chromatography High-purity waters 6.1.5.1 Thin-layer chromatography
436 437 437
6.2 Aromatic amines 6.2.1 Natural waters 6.2.1.1 Spectrophotometric method 6.2.1.2 Gas chromatography 6.2.1.3 High-performance liquid chromatography 6.2.1.4 Miscellaneous 6.2.1.5 Preconcentration 6.2.2 Sea water 6.2.2.1 High-performance liquid chromatography 6.2.3 Trade effluents 6.2.3.1 Spectrophotometric method 6.2.3.2 Gas chromatography 6.2.3.3 Miscellaneous 6.2.4 Waste waters 6.2.4.1 Spectrophotometric method 6.2.4.2 Gas chromatography 6.2.4.3 Miscellaneous
437 437 437 437 437 437 438 438 438 438 438 438 439 439 439 439 439
6.3 Chloroaromatic amines 6.3.1 Natural waters 6.3.1.1 Liquid chromatography 6.3.2 Waste waters 6.3.2.1 Gas chromatography
439 439 439 440 440
6.4 Aminophenols 6.4.1 Natural waters 6.4.1.1 Gas chromatography 6.4.2 Potable waters 6.4.2.1 High-performance liquid chromatography
440 440 440 440 440
6.5 Nitro compounds 6.5.1 Natural waters 6.5.1.1 Spectrophotometric method 6.5.1.2 High-performance liquid chromatography 6.5.2 Potable waters 6.5.2.1 High-performance liquid chromatography 6.5.2.2 Gas chromatography
440 440 440 440 440 440 441
Contents
6.5.3
6.5.4
Trade effluents 6.5.3.1 Spectrophotometric methods 6.5.3.2 Gas chromatography 6.5.3.3 Gas chromatography–mass spectrometry 6.5.3.4 Preconcentration Waste waters 6.5.4.1 High-performance liquid chromatography 6.5.4.2 Miscellaneous
xxi
441 441 441 441 442 442 442 442
6.6 Nitrophenols 6.6.1 Natural waters 6.6.1.1 Gas chromatography 6.6.1.2 Thin-layer chromatography 6.6.1.3 Column chromatography 6.6.1.4 Polarography
443 443 443 443 443 444
6.7 Nitriles 6.7.1 Natural waters 6.7.1.1 Titration and Spectrophotometric methods 6.7.1.2 Gas chromatography
444 444 444 444
6.8 Amides 6.8.1 Natural waters 6.8.1.1 Gas chromatography 6.8.1.2 High-performance liquid chromatography 6.8.1.3 Polarography 6.8.2 Sea water 6.8.2.1 High-performance liquid chromatography 6.8.3 Potable waters 6.8.3.1 Titration method 6.8.3.2 Spectrophotometric method 6.8.3.3 High-performance liquid chromatography 6.8.4 Trade effluents 6.8.4.1 Flow injection analysis 6.8.4.2 Gas chromatography 6.8.4.3 High-performance liquid chromatography 6.8.4.4 Size exclusion chromatography 6.8.4.5 Miscellaneous 6.8.5 Sewage effluents 6.8.5.1 High-performance liquid chromatography
444 444 444 445 445 445 445 446 446 446 446 446 446 446 446 446 447 447 447
xxii
Contents
6.9 Nitrosamines 6.9.1 Natural waters 6.9.1.1 Gas chromatography 6.9.1.2 Nuclear magnetic resonance spectrometry 6.9.2 Potable waters 6.9.2.1 Gas chromatography 6.9.3 Waste waters 6.9.3.1 Gas chromatography–mass spectrometry 6.9.4 Sewage effluents 6.9.4.1 Gas chromatography 6.9.5 Trade effluents 6.9.5.1 Gas chromatography 6.9.6 High-purity waters 6.9.6.1 Gas chromatography 6.9.6.2 Preconcentration
447 447 448 448 448 448 451 451 451 451 451 451 452 452 453
6.10 Amino acids 6.10.1 Natural waters 6.10.1.1 Gas chromatography 6.10.1.2 Fluorimetry 6.10.2 Sea and estuary waters 6.10.2.1 High-performance liquid chromatography 6.10.2.2 Fluorimetry 6.10.2.3 Preconcentration
453 453 453 453 453 453 455 455
6.11 Chloroamino acids 6.11.1 Waste waters 6.11.1.1 Gas chromatography–mass spectrometry
455 455 455
6.12 Nucleic acids 6.12.1 Natural waters 6.12.1.1 Spectrophotometric method 6.12.2 Sea water 6.12.2.1 Spectrophotometric method 6.12.2.2 Titration method
455 455 455 458 458 458
6.13 Ethylenediamine tetraacetic acid (EDTA) 6.13.1 Natural waters 6.13.1.1 Gas chromatography 6.13.1.2 Column chromatography 6.13.1.3 Polarography 6.13.1.4 Atomic absorption spectrometry
458 458 458 459 460 460
Contents
xxiii
6.13.2
Sea water 6.13.2.1 Polarography 6.13.3 Potable waters 6.13.3.1 Gas chromatography 6.13.4 Waste waters 6.13.4.1 High-performance liquid chromatography 6.13.5 Trade effluents 6.13.5.1 Liquid chromatography 6.13.6 Sewage effluents 6.13.6.1 Gas chromatography 6.13.6.2 Miscellaneous
460 460 461 461 461 461 461 461 461 461 462
6.14 Nitriloacitic acid 6.14.1 Natural waters 6.14.1.1 Spectrophotometric method 6.14.1.2 Gas chromatography 6.14.1.3 Polarography 6.14.1.4 Potential sweep potentiometry 6.14.1.5 Atomic absorption spectrometry 6.14.2 Sea water 465 6.14.2.1 Polarography 6.14.3 Potable waters 6.14.3.1 Gas chromatography 6.14.3.2 Fluorimetry 6.14.3.3 Polarography 6.14.4 Waste water 6.14.4.1 Titration method 6.14.4.2 Liquid chromatography 6.14.4.3 Atomic absorption spectrometry 6.14.4.4 Miscellaneous 6.14.5 Trade effluents 6.14.5.1 Liquid chromatography 6.14.6 Sewage effluents 6.14.6.1 Titration method 6.14.6.2 Gas chromatography 6.14.6.3 Ultraviolet spectrometry 6.14.6.4 Polarography 6.14.6.5 Miscellaneous
462 462 462 462 463 464 464
6.15 Other nitrogen compounds References
471 471
465 465 465 466 466 467 467 467 468 468 468 468 468 468 468 469 469 469
xxiv
7
Contents
Phosphorus-containing compounds
477
7.1 Alkyl and aryl phosphates 7.1.1 Natural waters 7.1.1.1 Gas chromatography 7.1.2 Estuary and sea waters 7.1.2.1 X-ray fluorescence spectrometry 7.1.3 Potable waters 7.1.3.1 Miscellaneous
477 477 477 478 478 478 478
7.2 Adenosine triphosphate 7.2.1 Natural waters 7.2.1.1 Luciferin–luciferase bioluminescence technique 7.2.1.2 Miscellaneous 7.2.2 Waste waters 7.2.2.1 Luciferin–luciferase bioluminescence technique 7.2.3 Sewage effluents 7.2.3.1 Luciferin–luciferase bioluminescence technique 7.2.4 Trade effluents 7.2.4.1 Luciferin–luciferase bioluminescence technique
478 479 479 480 480 480 480 480 481 481
7.3 Inositol phosphate esters 7.3.1 Natural waters 7.3.1.1 Miscellaneous
481 481 481
7.4 Organophosphorous compounds – general 7.4.1 Natural waters 7.4.1.1 Gas chromatographic detection with supported copper-cuprous oxide island film 7.4.1.2 Chemiresistant sensors for gas chromatographic detection 7.4.1.3 Surface acoustic wave sensors for gas chromatographic detection 7.4.1.4 Miscellaneous References
482 482
8
Sulphur-containing compounds
484
8.1 Mercaptans and disulphides 8.1.1 Natural waters 8.1.1.1 Titration method 8.1.1.2 Gas chromatography
484 484 484 484
482 482 482 482 482
Contents
8.1.2 8.1.3 8.1.4 8.1.5
Sea water 8.1.2.1 Gas chromatography Waste waters 8.1.3.1 Gas chromatography Trade effluents 8.1.4.1 Gas chromatography Sewage effluents 8.1.5.1 Gas chromatography 8.1.5.2 Gas chromatography–mass spectrometry 8.1.5.3 Miscellaneous
xxv
484 484 484 484 486 486 486 486 487 487
8.2 Benethionium salts 8.2.1 Waste waters 8.2.1.1 Spectrophotometric method
487 487 487
8.3 Hydroxymethanesulphinite salts 8.3.1 Trade effluents 8.3.1.1 Polarography
487 487 487
8.4 Benzothiazole, 2-mercaptobenzothiazole and 2-(4-morpholinyl)benzothiazole 8.4.1 Trade effluents 8.4.1.1 Gas chromatography–mass spectrometry 8.4.1.2 High-performance liquid chromatography
488 488 488 489
8.5 Dimethyl sulphoxide 8.5.1 Natural waters 8.5.1.1 Gas chromatography 8.5.1.2 Miscellaneous
489 489 490 492
8.6 Calcium lignosulphonate 8.6.1 Trade effluents 8.6.1.1 Gel permeation chromatography 8.6.1.2 Ultraviolet spectroscopy
493 493 493 493
8.7 Thiafluoron 8.7.1 Potable waters 8.7.1.1 High-performance liquid chromatography
493 493 493
8.8 Chlorolignosulphonic acids 8.8.1 Natural waters 8.8.1.1 Pyrolysis–gas chromatography–mass spectrometry with single-ion monitoring
493 493 493
xxvi
Contents
8.8.2 8.8.3
Potable waters 8.8.2.1 Pyrolysis–gas chromatography Trade effluents 8.8.3.1 Pyrolysis–gas chromatography
494 494 494 494
8.9 Tetrahydrothiophen 8.9.1 Natural waters 8.9.1.1 Gas chromatography
494 494 494
8.10 Organosulphur compounds, general References
494 494
9
496
Insecticides and herbicides
9.1 Chlorinated insecticides 9.1.1 Natural waters 9.1.1.1 Gas chromatography 9.1.1.2 Gas chromatography–mass spectrometry 9.1.1.3 Thin-layer chromatography 9.1.1.4 High-performance liquid chromatography 9.1.1.5 Nuclear magnetic resonance spectroscopy 9.1.1.6 Yeast hexokinase assay 9.1.1.7 Photodegradation technique 9.1.1.8 Miscellaneous 9.1.1.9 Preconcentration 9.1.1.10 Individual organochlorine insecticides 9.1.2 Aqueous precipitation 9.1.2.1 Gas chromatography 9.1.3 Sea water 9.1.3.1 Gas chromatography 9.1.3.2 Miscellaneous 9.1.3.3 Preconcentration 9.1.4 Potable waters 9.1.4.1 Gas chromatography 9.1.4.2 Thin-layer chromatography 9.1.5 Sewage effluents 9.1.5.1 Gas chromatography 9.1.5.2 Miscellaneous
496 496 496 510 510 512 512 513 513 513 514 515 519 519 519 519 521 522 522 522 523 524 524 524
9.2 Mixtures of chlorinated insecticides and polychlorobiphenyls 9.2.1 Natural waters 9.2.1.1 Gas chromatography
524 524 524
Contents
9.2.2
9.2.3 9.2.4 9.2.5
9.2.1.2 Supercritical fluid chromatography 9.2.1.3 Voltammetry 9.2.1.4 Miscellaneous 9.2.1.5 Preconcentration Sea water 9.2.2.1 Gas chromatography 9.2.2.2 Preconcentration Potable waters 9.2.3.1 Gas chromatography Waste waters 9.2.4.1 Gas chromatography Sewage effluents 9.2.5.1 Gas chromatography 9.2.5.2 Gas chromatography–mass spectrometry 9.2.5.3 Miscellaneous
9.3 Phosphorus-containing insecticides 9.3.1 Natural waters 9.3.1.1 Spectrophotometric methods 9.3.1.2 Flow injection analysis 9.3.1.3 Gas chromatography 9.3.1.4 Thin-layer chromatography 9.3.1.5 High-performance liquid chromatography 9.3.1.6 Reversed-phase adsorption liquid chromatography 9.3.1.7 Raman spectroscopy 9.3.1.8 Polarography 9.3.1.9 Enzymic assay methods 9.3.1.10 Miscellaneous 9.3.1.11 Preconcentration 9.3.1.12 Individual organophosphorous insecticides 9.3.2 Sea water 9.3.2.1 Spectrophotometric method 9.3.3 Potable waters 9.3.3.1 Gas chromatography 9.3.4 Waste waters 9.3.4.1 Gas chromatography 9.3.4.2 Field desorption mass spectrometry 9.3.4.3 High-performance liquid chromatography 9.3.5 Sewage effluents 9.3.5.1 Gas chromatography
xxvii
532 532 533 533 534 534 534 535 535 535 535 536 536 540 542 542 542 542 543 543 560 563 563 564 564 564 565 566 567 571 571 572 572 572 572 573 573 574 574
xxviii
Contents
9.3.5.2
Thin-layer chromatography
574
9.4 Pyrethrin insecticides 9.4.1 Natural waters 9.4.1.1 Gas chromatography
574 574 574
9.5 Thiram (tetramethylthiruam) insecticides 9.5.1 Natural waters 9.5.1.1 Spectrophotometric method 9.5.1.2 Voltammetry
575 575 575 575
9.6 Carbamate insecticides and herbicides 9.6.1 Natural waters 9.6.1.1 Spectrophotometric methods 9.6.1.2 Gas chromatography 9.6.1.3 Thin-layer chromatography 9.6.1.4 High-performance liquid chromatography 9.6.1.5 Enzymic assay 9.6.1.6 Fluorimetric analysis 9.6.1.7 Miscellaneous 9.6.1.8 Preconcentration 9.6.2 Waste waters 9.6.2.1 Gas chromatography 9.6.3 Trade effluents 9.6.3.1 High-performance liquid chromatography 9.6.4 Sewage effluents 9.6.4.1 High-performance liquid chromatography
575 575 575 576 580 581 581 581 582 582 582 582 582 582 582 582
9.7 Substituted urea-type herbicides 9.7.1 Natural waters 9.7.1.1 Gas chromatography 9.7.1.2 Thin-layer chromatography 9.7.1.3 High-performance liquid chromatography 9.7.1.4 Miscellaneous 9.7.1.5 Preconcentration
583 583 584 585 586 589 589
9.8 Triazine herbicides 9.8.1 Natural waters 9.8.1.1 Gas chromatography 9.8.1.2 Gas chromatography–mass spectrometry 9.8.1.3 Thin-layer chromatography 9.8.1.4 Enzymic immunoassay
589 589 590 591 592 592
Contents
9.8.1.5
9.8.2 9.8.3 9.8.4
High-resolution fast atom bombardment–mass spectrometry Miscellaneous
9.8.1.6 Sea water 9.8.2.1 Gas chromatography Waste water 9.8.3.1 Gas chromatography Sewage effluents 9.8.4.1 Gas chromatography–mass fragmentography
xxix
593 593 593 593 594 594 594 594
9.9 Phenoxyacetic acid herbicides 9.9.1 Natural waters 9.9.1.1 Gas chromatography 9.9.1.2 Thin-layer chromatography 9.9.1.3 Paper electrophoresis 9.9.1.4 High-performance liquid chromatography 9.9.1.5 Miscellaneous 9.9.2 Waste waters 9.9.2.1 Gas chromatography
594 595 595 598 598 599 599 601 601
9.10 Miscellaneous herbicides 9.10.1 Natural waters 9.10.2 Sea water 9.10.3 Potable waters 9.10.4 Waste waters
601 601 606 606 607
9.11 Growth regulators 9.11.1 Natural waters 9.11.1.1 Gas chromatography 9.11.1.2 Capillary isotachophoresis
607 607 607 608
9.12 Mixtures of pesticides and herbicides 9.12.1 Natural waters 9.12.1.1 Gas chromatography 9.12.1.2 Gas chromatography–mass spectrometry 9.12.1.3 Thin-layer chromatography 9.12.1.4 High-performance liquid chromatography 9.12.1.5 High-performance liquid chromatography– mass spectrometry 9.12.1.6 Mass spectrometry 9.12.1.7 Miscellaneous 9.12.1.8 Preconcentration References
608 608 608 609 610 611 612 612 612 612 613
xxx
Contents
10
Miscellaneous organic compounds
628
10.1 Squoxin piscicide 10.1.1 Natural waters
628 628
10.2 Coprostanol 10.2.1 Natural waters and sewage effluents
628 628
10.3 Fluorescent whitening agents 10.3.1 Natural waters
629 629
10.4 Mestranol and ethynyloestradiol 10.4.1 Natural waters
630 630
10.5 Carboxylmethyl succinate 10.5.1 Natural waters and sewage effluents 10.6 Isopropylmethylphosphonofluoridate (GB) and S-2(diisopropylamino)ethyl-O-ethylmethylphosphonothioate (VX) 10.6.1 Sea water
630 630
631 631
10.72,3-Dichloro-1,4-naphthaquinone 10.7.1 Natural waters
631 631
10.8 Geosmin and other water flavouring chemicals 10.8.1 Natural waters 10.8.2 Potable waters 10.8.2.1 Gas chromatography–mass spectrometry 10.8.2.2 Miscellaneous
631 632 632 632 633
10.9 Dihydroxyabietic acid 10.9.1 Trade effluents
633 633
10.10 Organic oxidants 10.10.1 Natural waters
633 633
10.11 Ozonization products 10.11.1 Natural waters 10.11.2 Potable waters 10.11.3 Waste waters 10.11.4 Sewage effluents 10.11.5 Swimming pool waters
633 633 634 636 636 638
10.12 Cobalamin (vitamin B12) 10.12.1 Natural waters
638 638
Contents
xxxi
10.12.2 Sea water 10.12.3 Sewage effluents
638 638
10.13 Humic and fulvic acids 10.13.1 Natural waters 10.13.1.1 Titration methods 10.13.1.2 Gel permeation chromatography 10.13.1.3 Ultraviolet spectroscopy 10.13.1.4 Fluorescence spectroscopy 10.13.1.5 Polarography 10.13.1.6 Miscellaneous 10.13.1.7 Preconcentration 10.13.2 Potable waters 10.13.2.1 Polarography 10.13.2.2 Miscellaneous 10.13.3 Sewage effluents
639 640 640 642 642 643 643 644 645 646 646 646 646
10.14 Chlorophyll and other plant pigments 10.14.1 Natural waters 10.14.1.1 Spectrophotometric methods 10.14.1.2 Thin-layer chromatography 10.14.1.3 High-performance liquid chromatography 10.14.1.4 Fluorescence spectroscopy 10.14.1.5 Miscellaneous 10.14.2 Sea water 10.14.2.1 Thin-layer chromatography 10.14.3 Sewage effluents 10.14.3.1 Miscellaneous References
648 648 648 650 651 653 655 656 656 658 658 660
11 Multiorganic mixtures in waters
664
11.1 Natural waters 11.1.1 Gas chromatography 11.1.2 Gas chromatography–mass spectrometry 11.1.3 High-performance liquid chromatography 11.1.4 Infrared spectroscopy 11.1.5 Raman spectroscopy 11.1.6 Supercritical fluid carbon dioxide extraction 11.1.7 Nuclear magnetic resonance spectroscopy 11.1.8 Miscellaneous 11.1.9 Preconcentration
664 664 664 667 672 673 673 675 675 677
xxxii
Contents
11.2 Potable waters 11.2.1 Gas chromatography–mass spectrometry 11.2.2 High-performance liquid chromatography–mass spectrometry 11.2.3 Reverse osmosis 11.2.4 Mass spectrometry 11.2.5 Miscellaneous 11.2.6 Preconcentration 11.2.6.1 Solvent extraction 11.2.6.2 Adsorption 11.2.6.3 Gas stripping 11.2.6.4 Headspace analysis
683 683
11.3 Rain water 11.3.1 Gas chromatography–mass spectrometry 11.3.2 High-performance liquid chromatography
697 697 697
11.4 Sea water 11.4.1 High-performance liquid chromatography 11.4.2 Ion-pair chromatography 11.4.3 Preconcentration
697 697 697 697
11.5 Waste waters 11.5.1 Gas chromatography 11.5.2 Gas chromatography–mass spectrometry 11.5.3 Headspace analysis 11.5.4 High-performance liquid chromatography 11.5.5 Nuclear magnetic resonance spectroscopy 11.5.6 Preconcentration 11.5.6.1 Gas chromatography 11.5.6.2 Headspace analysis 11.5.6.3 XAD-2 resin preconcentration
698 698 699 701 701 701 701 701 702 702
11.6 Trade effluents 11.6.1 Gas chromatography 11.6.2 Gas chromatography–mass spectrometry 11.6.3 Thin-layer chromatography 11.6.4 High-performance liquid chromatography 11.6.5 Infrared spectrometry 11.6.6 Miscellaneous 11.6.7 Preconcentration
702 702 702 706 706 706 708 708
683 689 690 690 690 690 692 695 695
Contents
xxxiii
11.7 Sewage effluents 11.7.1 Gas chromatography 11.7.2 Gas chromatography–mass spectrometry 11.7.3 High-performance liquid chromatography 11.7.4 Ion-exchange chromatography 11.7.5 Gel filtration chromatography 11.7.6 Preconcentration
708 708 709 710 711 711 711
11.8 Mineral waters 11.8.1 Preconcentration
712 712
11.9 Ultrapure waters 11.9.1 High-performance liquid chromatography References
712 712 712
12
722
Organometallic compounds
12.1 Organoarsenic compounds 12.1.1 Natural waters 12.1.1.1 Spectrophotometric methods 12.1.1.2 Gas chromatography 12.1.1.3 Ion-exchange chromatography 12.1.1.4 Polarography 12.1.1.5 Atomic absorption spectrometry 12.1.1.6 Miscellaneous 12.1.2 Sea water 12.1.2.1 Spectrophotometric method 12.1.2.2 Preconcentration 12.1.3 Potable waters 12.1.3.1 Spectrophotometric method 12.1.4 Waste waters 12.1.4.1 Spectrophotometric method
722 722 722 723 723 723 725 727 728 728 728 729 729 729 729
12.2 Organolead compounds 12.2.1 Natural waters 12.2.1.1 Gas chromatography 12.2.1.2 Polarography 12.2.1.3 Atomic absorption spectrometry 12.2.1.4 Miscellaneous 12.2.1.5 Preconcentration 12.2.2 Rain water 12.2.2.1 High-performance liquid chromatography
729 729 729 731 731 731 732 732 732
xxxiv
12.2.3
12.2.4 12.2.5
Contents
Waste waters 12.2.3.1 Spectrophotometric method 12.2.3.2 Atomic absorption spectrometry Trade effluents 12.2.4.1 Spectrophotometric method Potable waters 12.2.5.1 Gas chromatography–mass spectrometry
12.3 Organomercury compounds 12.3.1 Natural waters 12.3.1.1 Gas chromatography 12.3.1.2 Atomic absorption spectrometry 12.3.1.3 Inductively coupled plasma mass spectrometry 12.3.1.4 Photochroism-induced photoacoustic spectrometry 12.3.1.5 Neutron activation analysis 12.3.1.6 Miscellaneous 12.3.1.7 Preconcentration 12.3.2 Rain water 12.3.2.1 Atomic absorption spectrometry 12.3.3 Sea and coastal waters 12.3.3.1 Gas chromatography 12.3.3.2 Atomic absorption spectrometry 12.3.3.3 Miscellaneous 12.3.4 Potable waters 12.3.4.1 Atomic absorption spectrometry 12.3.4.2 Inductively coupled plasma mass spectrometry 12.3.4.3 Radiochemical method 12.3.4.4 Preconcentration 12.3.5 Waste waters 12.3.5.1 Spectrophotometric method 12.3.5.2 Atomic absorption spectrometry 12.3.6 Sewage effluents 12.3.6.1 Atomic absorption spectrometry 12.3.6.2 Thin-layer chromatography 12.3.6.3 Miscellaneous 12.3.7 Trade effluents 12.3.7.1 Spectrophotometric method 12.3.7.2 Gas chromatography 12.3.7.3 Atomic absorption spectrometry 12.3.7.4 Thin-layer chromatography
733 733 733 733 733 733 733 734 734 735 737 741 741 741 742 744 745 745 746 746 747 749 749 750 750 750 751 751 751 751 751 751 754 754 755 755 755 755 755
Contents
xxxv
12.4 Organotin compounds 12.4.1 Natural waters 12.4.1.1 Spectrophotometric method 12.4.1.2 Gas chromatography 12.4.1.3 Gas chromatography–mass spectrometry 12.4.1.4 Thin-layer chromatography 12.4.1.5 High-performance liquid chromatography 12.4.1.6 Spectrofluorimetry 12.4.1.7 Atomic absorption spectrometry 12.4.1.8 Miscellaneous 12.4.2 Rain water 12.4.2.1 Gas chromatography 12.4.3 Sea and estuary waters 12.4.3.1 Gas chromatography 12.4.3.2 High-performance liquid chromatography 12.4.3.3 Fluorescence spectrometry 12.4.3.4 Atomic absorption spectrometry 12.4.3.5 Miscellaneous 12.4.4 Potable waters 12.4.4.1 Gas chromatography 12.4.4.2 Spectrofluorimetry 12.4.5 Sewage effluents 12.4.5.1 Preconcentration
755 755 755 756 759 761 761 761 762 762 763 763 763 763 765 767 767 767 768 768 768 769 769
12.5 Organogermanium compounds 12.5.1 Natural waters 12.5.1.1 Atomic emission spectrometry
769 769 769
12.6 Organoantimony compounds 12.6.1 Natural waters 12.6.1.1 Atomic absorption spectrometry 12.6.1.2 Polarography
769 769 769 769
12.7 Organocopper compounds 12.7.1 Natural waters 12.7.1.1 High-performance liquid chromatography
770 770 770
12.8 Organocadmium compounds 12.8.1 Sea water 12.8.1.1 Anodic scanning voltammetry
770 770 770
12.9 Organosilicon compounds 12.9.1 Natural waters
770 770
xxxvi
Contents
12.9.1.1 12.9.1.2 12.9.1.3
Gas chromatography Mass spectrometry Inductively coupled plasma atomic emission spectrometry
770 770 771
12.10 Organoselenium compounds 12.10.1 Natural waters 12.10.1.1 Gas chromatography References
771 771 771 771
13
778
Determination of elements
13.1 Chlorine 13.1.1 Natural waters 13.1.1.1 Coulometric titration 13.1.1.2 Ion chromatography 13.1.1.3 Wickbold method 13.1.1.4 Neutron activation analysis 13.1.1.5 Miscellaneous 13.1.2 Potable waters 13.1.2.1 Coulometric titration 13.1.3 Waste waters 13.1.3.1 Titration method 13.1.4 Sewage effluents 13.1.4.1 Miscellaneous
778 778 778 778 779 779 781 781 781 781 781 783 783
13.2 Bromine 13.2.1 Natural waters 13.2.1.1 Ion chromatography 13.2.1.2 Neutron activation analysis 13.2.2 Potable waters 13.2.2.1 Coulometric titration 13.2.2.2 Neutron activation analysis
783 783 783 783 784 784 784
13.3 Iodine 13.3.1 Natural waters 13.3.1.1 Spectrophotometric methods 13.3.1.2 Neutron activation analysis 13.3.2 Potable waters 13.3.2.1 Neutron activation analysis 13.3.3 Waste waters 13.3.3.1 Titration method
784 784 784 784 785 785 785 785
Contents
xxxvii
13.4 Total halogens 13.4.1 Natural waters 13.4.1.1 Gas chromatography–mass spectrometry 13.4.2 Waste waters 13.4.2.1 Gas chromatography 13.4.2.2 Miscellaneous
785 785 785 785 785 786
13.5 Organic nitrogen 13.5.1 Natural waters 13.5.1.1 Kjeldahl digestion 13.5.1.2 Persulphate digestion 13.5.1.3 Spectrophotometric methods 13.5.1.4 Ultraviolet irradiation 13.5.1.5 Isotope dilution method 13.5.1.6 Miscellaneous 13.5.2 Sea water 13.5.2.1 Spectrophotometric method 13.5.3 Potable waters 13.5.3.1 Titration method 13.5.3.2 Kjeldahl digestion 13.5.4 Waste waters 13.5.4.1 Spectrophotometric method 13.5.4.2 Kjeldahl digestion 13.5.4.3 Miscellaneous 13.5.5 Trade effluents 13.5.5.1 Titration method 13.5.5.2 Kjeldahl digestion 13.5.5.3 Chemiluminescence analysis 13.5.6 Sewage effluents 13.5.6.1 Kjeldahl digestion 13.5.6.2 Flow injection analysis 13.5.6.3 Ammonia probe
786 786 786 787 787 787 790 793 793 793 795 795 795 795 795 795 796 796 796 796 796 797 797 797 797
13.6 Organic phosphorus 13.6.1 Natural waters 13.6.1.1 Digestion procedures 13.6.1.2 Spectrophotometric methods 13.6.1.3 Gas chromatography 13.6.1.4 Isotope exchange method 13.6.1.5 Miscellaneous
797 797 797 799 801 802 802
xxxviii
13.6.2
13.6.3
Contents
Waste waters 13.6.2.1 Spectrophotometric method 13.6.2.2 Inductively coupled plasma atomic emission spectrometry 13.6.2.3 Miscellaneous Trade effluents 13.6.3.1 Spectrophotometric method
803 803 803 803 803 803
13.7 Total organic phosphorus and total organic nitrogen 13.7.1 Natural waters 13.7.1.1 Digestion methods
804 804 804
13.8 Organosulphur compounds 13.8.1 Natural waters 13.8.1.1 Ion chromatography 13.8.1.2 Miscellaneous 13.8.2 Waste waters 13.8.2.1 Emission spectrometry
805 805 805 805 806 806
13.9 Organoboron compounds 13.9.1 Waste waters 13.9.1.1 Inductively coupled plasma atomic emission spectrometry 13.9.1.2 Spectrophotometric method 13.9.2 Sewage effluents 13.9.2.1 Fluorimetric method 13.9.3 Mineral waters 13.9.3.1 Ion-exchange chromatography 13.9.3.2 Spectrophotometric and spectrofluorimetric methods
806 806
13.10 Total organic carbon 13.10.1 Natural waters 13.10.1.1 Thermal combustion methods 13.10.1.2 Combustion method 13.10.1.3 Wet oxidation methods 13.10.1.4 Ultraviolet radiation decomposition procedures 13.10.1.5 Ultraviolet spectrometry 13.10.1.6 Electrochemical methods 13.10.1.7 Pyrolysis–gas chromatography–mass spectrometry
808 809 810 813 813
806 807 807 807 807 807 807
814 815 816 816
Contents
xxxix
13.10.1.8 Miscellaneous 13.10.2 Potable waters 13.10.2.1 Peroxydisulphate oxidation 13.10.2.2 Ultraviolet oxidation 13.10.2.3 Thermal combustion–ion chromatography 13.10.3 Waste waters 13.10.3.1 Miscellaneous 13.10.4 Trade effluents 13.10.4.1 Combustion methods 13.10.4.2 Reduction to methane 13.10.5 Sewage effluents 13.10.5.1 Combustion methods
817 818 818 819 819 820 820 821 821 822 822 822
13.11 Total inorganic carbon 13.11.1 Natural waters 13.11.1.1 Spectrophotometric method 13.11.1.2 Infrared analysis method 13.11.1.3 Miscellaneous
824 824 824 825 826
13.12 Dissolved organic carbon 13.12.1 Natural waters 13.12.1.1 Potassium persulphate–ultraviolet oxidation 13.12.1.2 Silver peroxydisulphate oxidation 13.12.1.3 Ultraviolet oxidation 13.12.1.4 Ultraviolet spectrometry 13.12.1.5 Miscellaneous 13.12.2 Potable waters 13.12.2.1 Miscellaneous 13.12.3 Trade effluents 13.12.3.1 Persulphate oxidation
826 826 826 827 827 828 828 829 829 829 829
13.13 Particulate organic carbon 13.13.1 Natural waters 13.13.1.1 Combustion methods 13.13.1.2 Miscellaneous 13.13.2 Rain water 13.13.2.1 Digestion method References
830 830 830 831 831 831 831
14
838
Oxygen demand parameters
14.1 Chemical oxygen demand 14.1.1 Natural waters
838 838
xl
Contents
14.1.2 14.1.3
14.1.4 14.1.5
14.1.1.1 Titration methods 14.1.1.2 Coulometric method 14.1.1.3 Spectrophotometric method Estuarine and sea waters 14.1.2.1 Miscellaneous Waste waters 14.1.3.1 Titration methods 14.1.3.2 Spectrophotometric methods 14.1.3.3 Flow injection analysis Trade effluents Sewage effluents 14.1.5.1 Titration methods
838 840 840 840 841 841 842 843 844 844 844 844
14.2 Total oxygen demand 14.2.1 Natural waters 14.2.1.1 Combustion methods 14.2.1.2 Miscellaneous 14.2.2 Waste waters 14.2.2.1 Miscellaneous 14.2.3 Trade effluents 14.2.4 Sewage effluents 14.2.4.1 Titration method 14.2.4.2 Electrochemical method 14.2.4.3 Miscellaneous
848 850 850 852 853 853 853 853 853 853 854
14.3 Biochemical oxygen demand 14.3.1 Natural waters 14.3.1.1 Automation of BOD procedure 14.3.1.2 Electrometric method 14.3.1.3 Effect of experimental parameters (temperature, partial pressure of oxygen, humidity) 14.3.1.4 Effect of light 14.3.1.5 Dilution water 14.3.1.6 Suppression of nitrification 14.3.1.7 Kinetics 14.3.2 Sea water 14.3.3 Waste waters 14.3.3.1 Microbial electrodes 14.3.3.2 Respirometric methods
854 855 856 856
857 857 858 858 858 859 860 863 863
Contents
xli
14.3.3.3
Correlations between oxygen demand parameters 14.3.4 Trade effluents 14.3.4.1 Relationship between oxygen demand parameters 14.3.5 Sewage effluents 14.3.5.1 Microbial electrodes 14.3.5.2 Kinetics References
867 868 870 870 871
Index
876
866 866
Preface
The presence of concentrations of organic substances in water is a matter of increasing concern to the water industry, environmentalists and the general public alike from the point of view of possible health hazards presented to both human and animal life, represented by domesticated and wild animals and bird and fish life. This awareness hinges on three facts: the increasing interest by the scientist and the public alike in matters environmental, an increased usage of organic materials in commerce coupled with the much wider variety of organic substances used nowadays, and finally, the availability of analytical methods sensitive enough to determine very low concentrations of these substances, the presence of which we formerly were unaware. It has been estimated that river waters can contain up to 2000 different organic substances over a wide concentration range and many of these survive processing in the water works and occur in potable water, with possible health implications. The Food and Drug Administration in America, amongst others, is systematically working its way through screening tests on these substances so far identified in water, but this is a process that will take many years to complete. As well as organics occurring in water as a direct result of industrial activity there are those which occur more indirectly from other causes, such as haloforms produced in the chlorination stage of the water treatment process, organometallic compounds produced by conversion of inorganic salts by biological activity in rivers and nitrosamine formation by conversion of inorganic nitrates. There are also, of course, naturally occurring organic substances in water. The purpose of this book is to draw together and systemize the body of information available throughout the world up to early 1998 on the occurrence and determination of organics of all types in non-saline and saline natural and treated water. In this way reference to a very scattered literature can be avoided. This is not a recipe book, i.e. methods are not presented in detail, space considerations alone would not permit this; instead the chemist is
Preface
xliii
presented with details of methods available for the determination of all types of organics in a variety of types of water samples. Methods are described in broad outline giving enough information for the chemist to decide whether he or she wishes to refer to the original paper. To this end, information is provided on applicability of methods, advantages and disadvantages of one method compared to another, interferences, sensitivity and detection limits. Examples of results obtained by various methods are given. Microbiological methods are not included as this subject would justify a separate book. Some enzymic assay methods are included. It was decided, on balance, to include organometallic compounds (Chapter 12) as these are now frequently being found in environmental water samples. Chemists wishing to identify unknown organic compounds in a sample increasingly employ combinations of chromatographic and mass spectrometric techniques. These techniques are discussed throughout the book and in more detail in Chapter 11. Where available, preconcentration techniques are discussed, enabling the sensitivity of methods to be improved by several orders of magnitude, a refinement often needed in environmental water analysis. Chapter 1, which forms an introduction, discusses the principles of the various techniques now being employed in water analysis, and the types of determinations to which these techniques can be applied. This chapter also contains a useful key system so that the reader can quickly locate in the book sections in which are discussed the determination by various techniques of particular organics in particular types of water sample. The contents are presented in as logical fashion as possible, starting in Chapter 2 with a discussion of hydrocarbons and polyaromatic hydrocarbons. Chapter 4 deals with the various types of surface active agents, whilst Chapters 3 and 5 – 8 deal, respectively, with oxygen, halogen, nitrogen, phosphorus and sulphur compounds. The determination of various types of insecticides and herbicides is discussed in Chapter 9. The book concludes with a discussion of elemental analysis and oxygen parameters (Chapters 13 and 14). Examination for organic substances combines all the exciting features of analytical chemistry. First, the analysis must be successfully completed and in many cases, such as spillages, must be completed quickly. Often the nature of the substances to be analysed for is unknown, the substances might occur at exceedingly low concentrations and might, indeed, be a complex mixture. To be successful in such an area requires analytical skills of a high order and the availability of sophisticated instrumentation. The work has been written with the interest of the following groups of people in mind: management and scientists in all aspects of the water
xliv
Preface
industry, river management, fishery industries, sewage effluent treatment and disposal, land drainage and water supply; also management and scientists in all branches of industry which produce aqueous effluents. It will also be of interest to agricultural chemists, agriculturists concerned with the ways in which organic chemicals used in crop or soil treatment permeate through the ecosystem, the biologists and scientists involved in fish, plant, insect and plant life, and also to the medical profession, toxicologists and public health workers and public analysts. Other groups or workers to whom the work will be of interest include oceanographers, environmentalists and, not least, members of the public who are concerned with the protection of our environment. Finally, it is hoped that the work will act as a spur to students of all subjects mentioned and assist them in the challenge that awaits them in ensuring that the pollution of the environment is controlled so as to ensure that by the turn of the century we are left with a worthwhile environment to protect. T.R. Crompton
Chapter 1
Introduction
1.1 Brief summary of methodologies 1.1.1 Titration procedures Anionic and cationic surface active agents, aliphatic amines, nitriles, amines, nucleic acid, nitriloacetic acid, mercaptans, disulphides, humic acid and fulvic acids, and also bromine, chlorine, iodine and organic nitrogen have been determined in a variety of water samples by titration procedures. As titration procedures are relatively insensitive compared to some other procedures, it is likely that they would be applied only to those types of water sample where the concentration of the determinand is relatively high, e.g. sewage and trade effluents, and not, for example, to potable water samples. The titration process has been automated so that batches of samples can be titrated non-manually and the data processed and reported via printouts and screens. One such instrument is the Metrohm 670 titroprocessor. This incorporates a built-in control unit and sample changer so that up to nine samples can be automatically titrated. The 670 titroprocessor offers incremental titrations with variable or constant volume steps (dynamic or monotonic titration). The measured value transfer in these titrations is either drift controlled (equilibrium titration) or effected after a fixed waiting time; pK determinations and fixed end points (e.g. for specified standard procedures) are naturally included. End-point titrations can also be carried out. Sixteen freely programmable computational formulae with assignment of the calculation parameters and units, mean-value calculations and arithmetic of one titration to another (via common variables) are available. Results can be calculated without any limitations. The 670 titroprocessor can also be used to solve complex analytical tasks. In addition to various auxiliary functions, which can be freely programmed, up to four different titrations can be performed on a single sample.
2
Introduction
In addition to the fully automated 670 system, Metrohm also supply simpler units with more limited facilities which nevertheless are suitable for more simple titrations. Thus the model 682 titroprocessor is recommended for routine titrations with automatic equivalence pointer cognition or to preset end points. The 686 titroprocessor is a lower-cost version of the above instrument, again with automatic equivalence point recognition and titration to preset end points. Mettler produce two automatic titrators, the DL 40 GP memotitrator and the lower-cost DL 20 compact titrator. Features available on the DL 40 GP include absolute and relative end-point titrations, equivalence-point titrations, back-titration techniques, multimethod applications, dual titration, pH stating, automatic learn titrations, automatic determination of standard deviation and means, series titrations, correction to printer, acid balance analogue output for recorder and correction to the laboratory information system. Up to 40 freely definable methods can be handled and up to 20 reagents held in store. Six control principles can be invoked. The DL 20 can carry out absolute (not relative) end-point titrations and equivalencepoint titrations, back-titration, series titrations, and correction to printer and balance and the laboratory information system. Only one freely definable method is available. Four control principles can be invoked. The DL 40 GP can handle potentiometric, voltammetric or photometric titrations. 1.1.2 Polarographic methods This technique has been applied to the following determinations in water, all of which are capable of undergoing an oxidation reduction (i.e. redox): polyaromatic hydrocarbons, aldehydes, amides, esters, lactams, quinones, anionic and non-ionic surface active agents, polychlorobiphenyls, chlorophenols, nitrophenols, ethylenediamine tetraacetic acid, nitriloacetic acid, hydroxymethylene sulphide salts, phosphoruscontaining insecticides, humic acid, fulvic acids and organocompounds of lead, antimony and arsenic. Differential pulse polarography has found limited applications (carboxylic acids and nitriloacetic acid), as has scanning voltammetry (chlorinated insecticides, polychlorobiphenols and organocadmium compounds). Three basic techniques of polarography are of interest and the basic principles of these are outlined below.
Brief summary of methodologies
3
Universal: differential pulse (DPN, DPI, DPR) In this technique a voltage pulse is superimposed on the voltage ramp during the last 40ms of controlled drop growth with the standard dropping mercury electrode; the drop surface is then constant. The pulse amplitude can be preselected. The current is measured by integration over a 20ms period immediately before the start of the pulse and again for 20ms as the pulse nears completion. The difference between the two current integrals (12 – 11) is recorded and this gives a peak-shaped curve. If the pulse amplitude is increased, the peak current value is raised but the peak is broadened at the same time. Classical direct current (DCT) In this direct current method, integration is performed over the last 20ms of the controlled drop growth (Tast procedure): during this time, the drop surface is constant in the case of the dropping mercury electrode. The resulting polarogram is step-shaped. Compared with classical DC polarography according to Heyrovsky, i.e. with the free-dropping mercury electrode, the DCT method offers great advantages: considerably shorter analysis times, no disturbance due to current oscillations, simpler evaluation and larger diffusion-controlled limiting current. Rapid square wave (SQW) Five square-wave oscillations of frequency around 125Hz are superimposed on the voltage ramp during the last 40ms of controlled drop growth – with the dropping mercury electrode the drop surface is then constant. The oscillation amplitude can be preselected. Measurements are performed in the second, third and fourth square-wave oscillation; the current is integrated over 2ms at the end of the first and at the end of the second half of each oscillation. The three differences of the six integrals (11 – 12, 13 – 14, 15 – 16) are averaged arithmetically and recorded as one current value. The resulting polarogram is peak shaped. Metrohm are leading suppliers of polarographic equipment. They supply three main pieces of equipment: the Metrohm 646 VA processor, the 647 VA stand (for single determinations) and the 675 VA sample changer for a series of determinations). Some features of the 646 VA processor are listed below: • • • •
Optimized data acquisition and data processing High-grade electronics for a better signal-to-noise ratio Automatic curve evaluation as well as automated standard addition for greater accuracy and smaller standard deviation Large, non-volatile methods memory for the library of fully developed analytical procedures
4
• • •
•
Introduction
Connection of the 675 VA sample changer for greater sample throughout Connection of an electronic balance Simple, perfectly clear operation principle via guidance in the dialogue mode yet at the same time high application flexibility thanks to the visual display and alphanumeric keyboard Complete and convenient result recording with built-in thermal recorder/printer
The 675 VA sample changer is controlled by the 646 VA processor on which the user enters the few control commands necessary. The 646 VA processor also controls the 677 drive unit and the 683 pumps. With these auxiliary units, the instrument combination becomes a polarographic analysis station which can be used to carry out on-line measurements. The 646 VA processor is conceived as a central, compact component for automated polarographic and voltammetric systems. Thus, two independent 647 VA stands or a 675 VA sample changer can be added. Up to four multidosimats of the 665 type for automated standard additions and/or addition of auxiliary solutions can be connected to each of these wet-chemical workstations. Connection of an electronic balance for direct transfer of data is also possible. Program-controlled automatic switching and mixing of these three electrode configurations during a single analysis via software commands occur. The complete electrode is pneumatically controlled. A hermetically sealed mercury reservoir of only a few millilitres suffices for approximately 200 000 drops. The mercury drops are small and stable; consequently, there is a good signal-to-noise ratio. Mercury comes into contact only with the purest inert gas and plastic free of metal traces. Filling is seldom required and very simple to carry out. The system uses glass capillaries which can be exchanged simply and rapidly. Up to 30 complete analytical methods (including all detailed information and instructions) can be filed in a non-volatile memory and called up. Consequently, a large extensive and correspondingly efficient library of analytical methods can be built up, comprehensive enough to carry out all routine determinations conveniently via call-up of a stored method. The standard addition method (SAM) is the procedure generally employed to calculate the analyte content from the signal of the sample solution. The SAM is coupled directly to the determination of the sample solution so that all factors which influence the measurement remain constant. There would be no doubt that the SAM provides results that have proved to be accurate and precise in virtually every case.
Brief summary of methodologies
5
The addition of standard solutions can be performed several times if need be (multiple standard addition) to raise the level of quality of the results still further. Normally, a real sample solution contains the substances to be analysed in widely different concentrations. In a single multi-element analysis, however, all components must be determined simultaneously. The superiority of the facilities offered by segmented data acquisition in this respect is clear when a comparison is made with previous solutions. The analytical conditions were inevitably a compromise; no matter what type of analytical conditions were selected, such large differences could rarely be reconciled. In the recording, either the peaks of some of the components were shown meaningfully – each of the other two were either no longer recognizable – or led to gigantic signals with cut-off peak tips. And all too often the differences were still too large even within the two concentration ranges. Since the recorder sensitivity and also all other instrument and electrode functions could only be set and adjusted for a single substance, even automatic range switching of the recorder was of very little use. The dilemma is solved with the 646VA processor: the freedom to divide the voltage sweep into substance-specific segments and to adjust all conditions individually and independently of one and another within these segments opens up quite new and, to date, unknown analytical possibilities. Furthermore, it allows optimum evaluation of the experimental data. Various suppliers of polarographs are summarized in Table 1.1. 1.1.3 Spectrophotometric methods 1.1.3.1 Visible spectrometry This technique is only of value when the identity of the compound to be determined is known. There are also limitations on the sensitivity that can be achieved, usually milligrams per litre or occasionally, micrograms per litre. The following types of compounds have been determined in water samples: aliphatic and aromatic hydrocarbons, carboxylic acids, alcohols, aldehydes, carbohydrates, dioxans, phenols, anionic, cationic and nonionic surface active agents, haloforms, chlorophenols, chlorinated isocyanurates, aliphatic and aromatic amines, nitro compounds, nitriles, amides, nucleic acids, nitriloacetic acid, benethionium salts, Thiram insecticide, carbamate insecticides and herbicides, chlorophyll and other plant pigments, organocompounds of arsenic, lead, mercury, tin,
6
Introduction
Table 1.1 Suppliers of polarographs
phosphorus-containing insecticides, carboxy methyl succinate, isopropylmethyl phosphonofluoridate, S–2 (disopropylamino)-ethyl-Oethylmethyl phosphonothioate and 2,3 dichloro-1,4-naphthaquinone. The technique has also been used to determine iodine, organic nitrogen and phosphorus, total inorganic carbon and chemical oxygen demand. Visible spectrophotometers are commonly used in the water industry for the estimation of colour in a sample or for the estimation of coloured products produced by reacting a colourless compound of the sample
Brief summary of methodologies
7
with a reagent, which produces a colour that can be evaluated spectrophotometrically. Some commercially available instruments, in addition to visible spectrophotometers, can also perform measurements in the UV and near IR regions of the spectrum. These have not yet found extensive application in the field of water analysis. Suppliers of visible spectrophotometers are reviewed in Table 1.2. 1.1.3.2 Ultraviolet spectrometry This technique has found limited application in water analysis: aliphatic, aromatic and polaromatic hydrocarbons, phenols, anionic and cationic surface active agents, saturated aliphatic halogen compounds, nitriloacetic acid and mimic and fulvic acids. The technique has also found application in the determination of total and dissolved organic carbon in water. 1.1.3.3 Luminescence and fluorescence spectrometry Spectrofluorimetric methods have been described for the determination of polyaromatic hydrocarbons, carboxylic acids, phenols, amino acids, polychlorobiphenyls, nitriloacetic acid, carbamate insecticides and herbicides, chlorophyll and other plant pigments, and organotin and organoboron compounds. Some applications of fluorescence spectrometry to the determination of polyaromatic hydrocarbons in water samples are summarized in Table 1.3. Generally speaking, concentrations down to the microgram per litre level can be determined by this technique with recovery efficiencies near 100%. Potentially, fluorimetry is valuable in every laboratory, including water laboratories, for the performance of chemical analysis where the prime requirements are selectivity and sensitivity. While only 5– 10% of all molecules possess a native fluorescence, many can be induced to fluoresce by chemical modification or being tagged with a fluorescent molecule. Luminescence is the generic name used to cover all forms of light emission other than that arising from elevated temperature (thermoluminescence). The emission of light through the absorption of UV or visible energy is called photoluminescence, and that caused by chemical reactions is called chemiluminescence. Light emission through the use of enzymes in living systems is called bioluminescence, the only known application of which to water analysis is the determination of adenosine triphosphate. Photoluminescence may be further subdivided into fluorescence, which is the immediate release (10-8s) of absorbed light
8
Introduction
Table 1.2 Visible–ultraviolet–near infrared spectrophotometers
Brief summary of methodologies
9
Table 1.3 Determination of polyaromatic hydrocarbons in water
energy, as opposed to phosphorescence, which is delayed release (10-6102s) of absorbed light energy. The excitation spectrum of a molecule is similar to its absorption spectrum, while the fluorescence and phosphorescence emissions occur at longer wavelengths than the absorbed light. The intensity of the emitted light allows quantitative measurement since, for dilute solutions, the emitted intensity is proportional to concentration. The excitation and emission spectra are characteristic of the molecule and allow qualitative measurements to be made. The inherent advantages of the technique, particularly fluorescence, are: 1 2 3
Sensitivity; picogram quantities of luminescent materials are frequently studied. Selectivity, derived from the two characteristic wavelengths. The variety of sampling methods that are available, i.e. dilute and concentrated samples, suspensions, solids, surfaces and combination with chromatographic methods, such as that used in the HPLC separation of o-phthalyl dialdehyde derivatized amino acids in natural and sea water samples.
10
Introduction
Fluorescence spectrometry forms the majority of luminescence analysis. However, the recent developments in instrumentation and room-temperature phosphorescence techniques have given rise to practical and fundamental advances which should increase the use of phosphorescence spectrometry. The sensitivity of phosphorescence is comparable to that of fluorescence and complements the latter by offering a wider range of molecules of study. The pulsed xenon lamp forms the basis for both fluorescence and phosphorescence measurement. The lamp has a pulse duration at halfpeak height of 10µs. Fluorescence is measured at the instant of the flash. Phosphorescence is measured by delaying the time of measurement until the pulse has decayed to zero. Several methods are employed to allow the observation of phosphorescence. One of the most common techniques is to supercool solutions to a rigid glass state, usually at the temperature of liquid nitrogen (77K). At these temperatures molecular collisions are greatly reduced and strong phosphorescence signals are observed. Under certain conditions phosphorescence can be observed at room temperature from organic molecules adsorbed on solid supports such as filter paper, silica and other chromatographic supports. Phosphorescence can also be detected when the phosphor is incorporated into an ionic micelle. Deoxygenation is still required either by degassing with nitrogen or by the addition on sodium sulphite. Micellestabilized room-temperature phosphorescence (MS RTP) promises to be a useful analytical tool for determining a wide variety of compounds such as pesticides and polyaromic hydrocarbons. Perkin–Elmer and Hamilton both supply luminescence instruments.
Perkin–Elmer LS-3B and LS-5B luminescence spectrometers The LS-3B is a fluorescence spectrometer with separate scanning monochromators for excitation and emission, and digital displays of both monochromator wavelengths and signal intensity. The LS-5B is a ratioing luminescence spectrometer with the capability of measuring fluorescence, phosphorescence and bio- and chemiluminescence. Both instruments are equipped with a xenon discharge lamp source and have an excitation wavelength range of 230–720nm and an emission wavelength range of 250–800nm. These instruments feature keyboard entry of instrument parameters which, combined with digital displays, simplifies instrument operation. A high-output pulsed xenon lamp, having low power consumption and minimal ozone production, is incorporated within the optical module.
Brief summary of methodologies
11
Through the use of an RS 232C interface, both instruments may be connected to Perkin–Elmer computers for instrument control and external data manipulation. With the LS-5B instrument, the printing of the sample photomultiplier can be delayed so that it no longer coincides with the flash. When used in this mode, the instrument measures phosphorescence signals. Both the delay of the start of the gate (t ) and the duration of the gate (t ) can be d g selected in multiples of 10µs from the keyboard. This corrects excitation and emission spectra. Delay times may be accurately measured by varying the delay time and noting the intensity at each value. Specificity in luminescence spectrometry is achieved because each compound is characterized by an excitation and emission wavelength. The identification of individual compounds is made difficult in complex mixtures because of the lack of structure from conventional excitation or emission spectra. However, by collecting emission and excitation spectra for each increment of the other, a fingerprint of the mixture can be obtained. This is visualized in the form of a time-dimensional contour plot on a three-dimensional isometric plot. Fluorescence spectrometers are equivalent in their performance to single-beam UV–visible spectrometers in that the spectra they produce are affected by solvent background and the optical characteristics of the instrument. These effects can be overcome by using software built into the Perkin–Elmer LS-5B instrument or by using application software for use with the Perkin–Elmer 3700 and 7700 computers. Perkin–Elmer LS-2B microf ilter f luorimiter The model LS-2B is a low-cost, easy-to-operate, filter fluorimeter that scans emission spectra over the wavelength range 390–700nm (scanning) or 220–650nm (individual interference filters). The essentials of a filter fluorimeter are as follows: • • • •
a source of UV/visible energy (pulsed xenon); a method of isolating the excitation wavelength; a means of discriminating between fluorescence emission and excitation energy; a sensitive detector and a display of the fluorescence intensity.
The model LS-2B has all these features arranged to optimize sensitivity for microsamples. It can also be connected to a highly sensitive 7µL liquid chromatographic detector for detecting the constituents in the column effluent. It has the capability of measuring fluorescence, time-resolved fluorescence, and bio- and chemiluminescent
12
Introduction
signals. A 40-portion autosampler is provided. An excitation filter kit containing six filters – 310, 340, 375, 400, 450 and 480nm – is available. 1.1.3.4 Infrared and Raman spectrometry Both these techniques have only limited application to the analysis of water samples: • •
Infrared spectrometry: aliphatic and aromatic hydrocarbons; anionic surface active agents; humic and fulvic acids, total inorganic carbon. Raman spectrometry: polyaromatic hydrocarbons, phenols; calcium lignosulphate, phosphorus-containing insecticides.
In addition, both techniques have been applied to the analysis of mixtures of different typical organic compounds in water. A more recent development is Fourier transform infrared analysis. Fourier transform infrared spectrometry Fourier transform infrared spectrometry, a versatile and widely used analytical technique, relies on the creation of interference in a beam of light. A source light beam is split into two parts and a continually varying phase difference is introduced into one of the resultant beams. The two beams are recombined and the interference signal is measured and recorded as an interferogram. A Fourier transform of the interferogram provides the spectrum of the detected light. Fourier transform infrared spectrometry, a seemingly indirect method of spectrometry, has many practical advantages, as discussed below. A Fourier transform infrared spectrometer consists of an infrared source, an interference modulator (usually a scanning Michelson interferometer), a sample chamber and an infrared detector. Interference signals measured at the detector are usually amplified and then digitized. A digital computer initially records and then processes the interferogram and also allows the spectral data that results to be manipulated. Permanent records of spectral data are created using a plotter or other peripheral device. The principal reasons for choosing Fourier transform infrared spectrometers are first, that these instruments record all wavelengths simultaneously and thus operate with maximum efficiency and second, that they have a more convenient optical geometry than do dispersive infrared instruments. These two facts lead to the following advantages: •
Fourier transform infrared spectrometers achieve much higher signalto-noise ratios in comparable scanning times.
Brief summary of methodologies
•
• • •
13
They can cover wide spectral ranges with a single scan in a short scan time, thereby permitting the possibility of kinetic time-resolved measurements. They provide higher-resolution capabilities without undue sacrifices in energy throughput or signal-to-noise ratios. They encounter none of the stray light problems usually associated with dispersive spectrometers. They provide a more convenient beam geometry – circular rather than slit shaped – at the sample focus.
Conventional Raman spectroscopy cannot be applied directly to water samples, although it is occasionally used to provide information on organic solvent extracts of water samples. Fourier transform Raman spectroscopy, on the other hand, can be directly applied to water samples. The technique complements infrared spectroscopy in that some functional groups (e.g. unsaturation) give a much stronger response in the infrared. Several manufacturers (Perkin–Elmer, Digilab, Bruker) now supply Fourier transform infrared spectrometers. 1.1.4 Flow injection analysis This technique has found limited application in water analysis and is particularly useful when routine automated analysis at the microgram per litre level of large numbers of water samples is required. Determinations of the following determinands have been described: phenols, cationic and non-ionic surface active agents, amides, phosphorus-containing insecticides, also organic nitrogen and chemical oxygen demand. Flow-injection analysis (FIA) is a rapidly growing analytical technique. Since the introduction of the original concept by Ruzicka and Hansen [13] in 1975, about 1000 papers have been published. Flow-injection analysis is based on the introduction of a defined volume of sample into a carrier (or reagent) stream. This results in a sample plug bracketed by carrier (Fig. 1.1 (a)). The carrier stream is merged with a reagent stream to obtain a chemical reaction between the sample and the reagent. The total stream then flows through a detector (Fig. 1.1 (b)). Although spectrophotometry is the commonly used detector system in this application, other types of detectors have been used, namely fluorimetric, atomic absorption emission spectrometry and electrochemical (e.g. ion selective electrodes). The pump provides constant flow and no compressible air segments are present in the system. As a result the residence time of the sample in the system is absolutely constant. As it moves towards the detector the sample is mixed with both carrier and reagent. The degree of dispersion
14
Introduction
Fig. 1.1 (a) Schematic diagram of the flow pattern in an FIA system directly after injection of sample, (b) Simple FIA system for one reagent; S denotes the sample injection site and D is the flow-through detector, (c) Typical FIA peaks (detector output signals), (d) Radial and axial dispersion in an injected sample plug, (e) Rapid scan of an FIA curve, (f) Configuration of an FIA system.
(or dilution) of the sample can be controlled by varying a number of factors, such as sample volume, length and diameter of mixing coils and flow rates. When the dispersed sample zone reaches the detector, neither the chemical reaction nor the dispersion process has reached a steady state. However, experimental conditions are held identical for both samples and standards in terms of constant residence time, constant temperature and constant dispersion. The sample concentration can thus be evaluated against appropriate standards injected in the same manner as samples (Fig. 1.1(c)).
Brief summary of methodologies
15
The short distance between the injection site and the merging point ensures negligible dispersion of the sample in this part of the system. This means that sample and reagent are mixed in equal proportions at the merging point. The mixing technique can be best understood by having a closer look at the hydrodynamic conditions in and around the merging point (Fig. 1.1(d)). In Fig. 1.1 (d) the hydrodynamic behaviour is simplified in order to explain the mixing process. Let us assume that there is no axial dispersion and that radial dispersion is complete when the sampler reaches the detector. The volume of the sample zone is thus 200µg after the merging point (100µL sample + 100µL reagent as flow rates are equal). The total flow rate is 2.0mL min-1. Simple mathematics then gives a residence time of 6s for the sample in the detector flow cell. In reality, response curves reflect some axial dispersion. A rapid scan curve is shown in Fig. 1.1 (e). The baseline is reached within 20s. This makes it possible to run three samples per minute and obtain baseline readings between each sample (no carryover), i.e. 180 samples per hour. The configuration of an FIA system is shown schematically in Fig. 1.1(f). The (degassed) carrier and reagent solution(s) must be transported in a pulse-free transport system and at a constant rate through narrow Teflon (Du Pont) tubing. In a practical FIA system, peristaltic pumps are usually used since they have several channels, and different flow rates may be achieved by selection of a pump tube with a suitable inner diameter. A manifold provides the means of bringing together the fluid lines and allowing rinsing and chemical reaction to take place in a controlled way. Manifolds with several lines can be assembled as required. These manifolds are mounted on plastic trays and allow the use of different reaction coils. Flow-injection analysers available range from relatively low-cost unsophisticated instruments such as those supplied by Advanced Medical Supplies, Skalar and Chemlab to the very sophisticated instruments such as the FIA star 5010 and 5020 supplied by Tecator (Table 1.4). 1.1.5 A tomic spectrometric methods 1.1.5.1 Atomic absorption spectrometry Basically, the atomic absorption method was designed for the determination of cations. However, it has been applied to the indirect determination of some organic substances. If an excess of a metal ion is
16
Introduction
Table 1.4 Equipment for flow-injection analysis
Brief summary of methodologies
17
added to a sample containing an organic substance which complexes with that metal and then the complex is extracted from the sample with a suitable organic solvent, then determination of the uncomplexed excess metal in the water phase enables one to estimate the amount of metal that has been complexed and therefore the concentration of the organic substance; alternatively, the complexed metal content of the organic extract can be determined. Thus, anionic surface active agents form a chloroform soluble complex with bis(ethylenediamine) copper (II) ion. Determination of copper in the chloroform extract enables one to estimate the concentration of anionic in the original water sample. Substances that have been determined by this technique include anionic and non-ionic surface active agents, ethylenediamine tetraacetic acid, nitriloacetic acid, and organic compounds of arsenic, lead, mercury, tin and antimony. Since shortly after its inception in 1955, atomic absorption spectrometry has been the standard tool employed by analysts for the determination of trace levels of metals in water samples. In this technique a fine spray of the analyte is passed into a suitable flame, frequently oxygen, acetylene or nitrous oxide acetylene, which converts the elements to an atomic vapour. Through this vapour radiation is passed at the right wavelength to excite the ground state atoms to the first excited electronic level. The amount of radiation absorbed can then be measured and directly related to the atom concentration: a hollow cathode lamp is used to emit light with the characteristic narrow line spectrum of the analyte element. The detection system consists of a monochromator (to reject other lines produced by the lamp and background flame radiation) and a photomultiplier. Another key feature of the technique involves modulation of the source radiation so that it can be detected against the strong flame and sample emission radiation. A limitation of this technique is its lack of sensitivity compared to that available by other techniques (e.g. inductively coupled plasma atomic emission spectrometry, Section 1.1.5.2). Suitable instrumentation is listed in Table 1.5. 1.1.5.2 Inductively coupled plasma atomic emission spectrometry This technique has, in recent years, been found to be particularly useful for the determination in water of extremely low levels of organic compounds of mercury, silicon, phosphorus and boron. An inductively coupled plasma is formed by coupling the energy from a radiofrequency (1–3kW or 27–50MHz) magnetic field to free electrons in a suitable gas. The magnetic field is produced by a two- or three-turn water-cooled coil and the electrons are accelerated in circular paths around the magnetic field lines that run axially through the coil. The
Table 1.5 Available flame and graphite furnace atomic absorption spectrometers
20
Introduction
initial electron ‘seeding’ is produced by a spark discharge but, once the electrons reach the ionization potential of the support gas, further ionization occurs and a stable plasma is formed. The neutral particles are heated indirectly by collisions with the charged particles upon which the field acts. Macroscopically the process is equivalent to heating a conductor by a radio-frequency field, the resistance to eddy-current flow producing Joule heating. The field does not penetrate the conductor uniformly and therefore the largest current flow is at the periphery of the plasma. This is the so-called ‘skin’ effect and, coupled with a suitable gas-flow geometry, it produces an annular or doughnut-shaped plasma. Electrically, the coil and plasma form a transformer with the plasma acting as a one-turn coil of finite resistance. The properties of an inductively coupled plasma closely approach those of an ideal source for the following reasons: •
•
• •
•
•
The source must be able to accept a reasonable input flux of the sample and it should be able to accommodate samples in the gas, liquid or solid phases. The introduction of the sample should not radically alter the internal energy generation process or affect the coupling of energy to the source from external supplies. The source should be operable on commonly available gases and should be available at a price that will give cost-effective analysis. The temperature and residence time of the sample within the source should be such that all the sample material is converted to free atoms irrespective of its initial phase or chemical composition; such a source should be suitable for atomic absorption or atomic fluorescence spectrometry. If the source is to be used for emission spectrometry, then the temperature should be sufficient to provide efficient excitation of a majority of elements in the periodic table. The continuum emission from the source should be of a low intensity to enable the detection and measurement of weak spectral lines superimposed upon it, the sample should experience a uniform temperature field and the optical density of the source should be low so that a linear relationship between the spectral line intensity and the analyte concentration can be obtained over a wide concentration range.
Greenfield et al. [14] were the first to recognize the analytical potential of the annular inductively coupled plasma. Wendt and Fassel [15] reported early experiments with a ‘tear drop’-shaped inductively coupled plasma
Brief summary of methodologies
21
but later described the medium-power (1–3kW), 18mm annular plasma now favoured in modern analytical instruments [16]. The current generation of inductively coupled plasma emission spectrometers provide limits of detection in the range 0.1–500µg L-1 in solution, a substantial degree of freedom from interference and a capability for simultaneous multi-element determination facilitated by a directly proportional response between the signal and the concentration of the analyte over a range of about five orders of magnitude. The most common method of introducing liquid samples into the inductively coupled plasma is by using pneumatic nebulization [24] in which the liquid is dispensed into a fine aerosol by the action of a high-velocity gas stream. To allow the correct penetration of the central channel of the inductively coupled plasma by the sample aerosol, an injection velocity of about 7m s -1 is required. This is achieved using a gas injection with a flow rate of about 0.5–11 min-1 through an injector tube of 1.5–2.0mm internal diameter. Given that the normal sample uptake is 1–2mL min -1 this is an insufficient quantity of gas to produce efficient nebulization and aerosol transport. Indeed, only 2% of the sample reaches the plasma. The fine gas jets and liquid capillaries used in inductively coupled plasma nebulizers may cause inconsistent operation and even blockage when solutions containing high levels of dissolved solids, such as sea water and particulate matter, are used. Such problems have led to the development of a new type of nebulizer, the most successful being based on a principle originally described by Babington (US Patents). In these, the liquid is pumped from a widebore tube and thence conducted to the nebulizing orifice by a V-shaped groove [17] or by the divergent wall of an overexpanded nozzle [18]. Such devices handle most liquids and even slurries without difficulty. Nebulization is inefficient and therefore not appropriate for very small liquid samples. Introducing samples into the plasma in liquid form reduces the potential sensitivity because the analyte flux is limited by the amount of solvent that the plasma will tolerate. To circumvent these problems a variety of thermal and electrothermal vaporization devices have been investigated. Two basic approaches are in use. The first involves indirect vaporization of the sample in an electrothermal vaporizer, e.g. a carbon rod or tube furnace or heated metal filament as commonly used in atomic absorption spectrometry [19–21]. The second involves inserting the sample into the base of the inductively coupled plasma on a carbon rod or metal filament support [22, 23]. Available instrumentation is reviewed in Table 1.6.
Table 1.6 Inductively coupled plasma optical emission spectrometers available on the market
24
Introduction
1.1.5.3 Atomic emission spectrometry This technique has been used only for the determination of sulphur- and germanium-containing organic compounds in water. 1.1.6 Nuclear magnetic resonance spectrometry This technique has found limited application in water analysis, namely with nitrosamines, chlorinated insecticides, humic and fulvic acids and mixtures of organic compounds. Instrument suppliers: • •
Gemini Superconducting Fourier transform, NMR systems, VXR series 5: Varian Instruments, Sugar Lane, Texas, USA. NMR imaging spectrometer systems: Vis, 1120 Auburn Road, Fremont, California 94538, USA.
1.1.7 Neutron activation analysis This is a very sensitive technique whose application in water has been limited to the determination of chlorine, total chlorine, total iodine and organomercury compounds. Due to the complexity and cost of the technique, no water laboratory in the UK has its own facility for carrying out neutron activation analysis. Instead, samples are sent to one of the organizations that possess the facilities, e.g. the Atomic Energy Research Establishment at Harwell or the Joint Manchester-Liverpool University Reactor located at Risley. The technique’s extreme sensitivity makes it suitable for use when a referee analysis is required on a material which has become a standard for checking out other methods. Another advantage of the technique is that a foreknowledge of the elements present is not essential. It can be used to indicate the presence and concentration of entirely unexpected elements, even when present at very low concentrations. In neutron activation analysis, the sample in a suitable container, often a pure polyethylene tube, is bombarded with slow neutrons for a fixed time together with standards. Transmutations convert analyte elements into radioactive elements, which are either different elements or isotopes of the original analyte. After removal from the reactor the product is subject to various counting techniques and various forms of spectrometry to identify the elements present and their concentration.
Brief summary of methodologies
25
1.1.8 X-ray fluorescence spectrometry The only application of this technique in water analysis discussed in the literature to date is the determination of alkyl and aryl phosphate esters. Due to its true multi-element analysis capability it is possible that further elemental applications to water analysis may be found, such as analysis for total organically bound chlorine, sulphur, phosphorus and silicon. 1.1.8.1 Energy-dispersive X-ray fluorescence spectrometry Energy-dispersive X-ray fluorescence (EDXRF) spectrometry is an instrumental analytical technique for non-destructive multi-elemental analysis. The use of modern-day technologies coupled with the intrinsic simplicity of X-ray fluorescence spectra (as compared for instance with optical emission (OE) spectra) means that the powerful EDXRF technique can be used routinely. The EDXRF spectrum for iron is a clearly resolved doublet, whilst the optical emission spectrum contains more than 4000 lines. This simplicity is a direct consequence of the fact that XRF spectra are a result of inner shell electron transitions which are possible only between a limited number of energy levels for the relatively few electrons. Optical emission spectra, on the other hand, arise from electron transitions in the outer, valence shells which are closer together in energy, more populated than the inner shells and from which it is easier to promote electron transitions. In order to generate X-ray spectra, we may excite the elements in the specimen with any one of the following: • • • • •
X-ray photons high-energy electrons high-energy charged particles gamma rays synchrotron radiation
The term XRF is generally applied when X-ray photons are used to generate characteristic X-rays from the elements in the specimen. The most commonly used source of such X-rays (in the 2–100keV range) are radioisotopes and X-ray tubes. An EDXRF spectrometer such as the XR300 uses a compact, low-power (10–100W typical) X-ray tube capable of delivery of X-ray photons with a maximum energy of 30 or 50keV.
26
Introduction
Why is the technique referred to as ‘energy-dispersive’ XRF? The classical XRF spectrometer, which has been commercially available since the 1950s uses crystal structures to separate (resolve) the X-rays emanating from the fluorescence process in the irradiated specimen. These crystals diffract the characteristic X-rays from the elements in the specimen, allowing them to be separated and measured. The characteristic fluorescent X-rays are said to have been separated from each other by the process of ‘wavelength dispersion’ (WDXRF). Each element emits characteristic lines which can be separated by WDXRF before being individually counted. For each line and diffracting crystal, we can set a detector at a particular angle (from the Bragg equation) and collect X-rays, which are primarily from the selected element. The EDXRF system uses the Si(Li) (lithium-drifted silicon) detector to simultaneously collect all X-ray energies emitted from the specimen. Each detected X-ray photon gives rise to a signal from the detector. The magnitude of this signal is proportional to the energy of the detected X-ray and when amplified and digitized can be passed to a multichannel analyser which displays a histogram of number of X-rays (intensity) against energy. The incident photons, therefore, have been electronically separated (dispersed) according to their energy. The energy of each of the X-rays from all the elements is readily accessible from published tables. Due to the simple spectra and the extensive element range (sodium upwards) which can be covered using the Si(Li) detector and a 50kV Xray tube, EDXRF spectrometry is perhaps unparalleled for its quantitative element analysis power. Qualitative analysis is greatly simplified by the presence of few peaks which occur in predictable positions and by the use of tabulated element/line markers which are routinely available from the computerbased analyser. To date, the most successful method of combined background correction and peak deconvolution is to use the method of digital filtering and least squares (FLS) fitting of reference peaks to the unknown spectrum [25]. This method is robust, simple to automate and is applicable to any sample type. The combination of the digital filtering and least squares peak deconvolution method and empirical correction procedures has application throughout elemental analysis. This approach is suitable for specimens of all physical types and is used in a wide selection of industrial applications.
Brief summary of methodologies
27
1.1.8.2 Total reflection X-ray fluorescence spectrometry The major disadvantage of conventional energy dispersive X-ray fluorescence spectrometry has been poor elemental sensitivity, a consequence of high background noise levels resulting mainly from instrumental geometries and sample matrix effects. Total reflection X-ray fluorescence (TXRF) is a relatively new multi-element technique with the potential to be an impressive analytical tool for trace-elemental determinations for a variety of sample types. The fundamental advantage of TXRF is its capability to detect elements in the picogram range in comparison to the nanogram levels typically achieved by traditional energy-dispersive X-ray fluorescence spectrometry. The problem in detecting atoms at the nanogram per litre or submicrogram per litre level is basically one of being able to obtain a signal which can be clearly distinguished from the background. The detection limit being given typically as the signal which is equivalent to three times the standard deviation of the background counts for a given unit of time. In energy-dispersive X-ray fluorescence spectrometry the background is essentially caused by interactions of radiation with matter resulting from an intense flux of elastic and Compton-scattered photons. The background, especially in the low-energy region (0–20keV), is due in the main to Compton scattering of high-energy Bremsstrahlung photons from the detector crystal itself. In addition, impurities on the specimen support will contribute to the background. The Auger effect does not contribute to an increased background, as the emitted electrons, of different but low energy, are absorbed either in the beryllium foil of the detector entrance windows or in the air path of the spectrometer. A reduction in the spectral background can be effectively achieved by X-ray total reflection at the surface of a smooth reflector material such as quartz. X-ray total reflection occurs when an X-ray beam impinges on a surface at less than the critical angle of total reflection. If a collimated Xray beam impinges onto the surface of a plane smooth and polished reflector at an angle less than the critical angle, then total reflection occurs. In this case the angle of incidence is equal to the angle of reflection and the intensities of the incident and totally reflected beams should be equal. The principles of TXRF were first reported by Yaneda and Horiuchi [26] and further developed by Aiginger and Wodbrauschek [27]. In TXRF the existing primary X-ray beam impinges upon the specimen prepared as a thin film on an optically flat support at angles of incidence in the region of 2–5 minutes of arc below the critical angle. In practice the primary radiation does not (effectively) enter the surface of the support but skims the surface, irradiating any sample placed on the support surface. The scattered radiation from the sample support is
28
Introduction
virtually eliminated, thereby drastically reducing the background noise. A further advantage of the TXRF system, resulting from the new geometry used, is that the solid-state energy-dispersive detector can be accommodated very close to the sample (0.3mm), which allows a large solid angle of fluorescent X-ray collection, thus enhancing signal sensitivity and enabling the analysis to be carried out in air at atmospheric pressure. The sample support or reflector is a 3cm diameter wafer made of synthetic quartz or Perspex. The water sample can be placed directly onto the surface. The simplest way to prepare liquid samples is to pipette volumes between 1 and 50µL directly onto a quartz reflector and allow them to dry. For aqueous solutions the reflector can be made hydrophobic (e.g. by silicon treatment) in order to hold the sample in the centre of the plate. Suitable elements for calibration can be achieved by a simple standard addition technique. Since Yaneda and Horiuchi [26] first reported the use of TXRF various versions have been developed (Knoth et al. [28, 29]; Schwenke and Knoth [30]; Pella and Dobbyn [31]. Recently an X-ray generator with a fine focus tube and multiple reflection optics has been developed by Seifert & Co. and coupled with an energy-dispersive spectrometer fitted with an Si(Li) detector and multi-channel analyser supplied by Link Analytical. The new system, which will be described later, known as the EXTRA II, represents the first commercially available TXRF instrument. An example of the detection limits achieved by the Link Analytical EXTRA II (3s above background, counting time 1000s) are shown in Fig. 1.2 for the molybdenum anode X-ray tube and for excitation with the filtered Bremsstrahlung spectrum for a tungsten X-ray tube. The data shown was obtained with diluted aqueous solutions which can be considered to be virtually free from any matrix effects. A detection limit of 10pg for the 10µL sample corresponds to a concentration of 1µg L-1. A linear dynamic range of four orders of magnitude is obtained for most elements. The attractive features of TXRF can be summarized as follows: • • • •
An inherent universal calibration curve is obtained as a smooth function of atomic number. The use of internal single-element standardization eliminates the need for matrix-matched external standards. Only small sample volumes are required (5–50µL). The technique requires only a simple sample preparation methodology.
The attractive features of the TXRF technique outlined above suggest that TXRF has the potential to become a very powerful analytical tool for
Brief summary of methodologies
29
Fig. 1.2 Minimum detection limits of the TXRF spectrometer (Mo-tube 13mA/60kV; counting time 1000s).
trace-elemental determinations applicable to a wide range of matrix types and may, indeed, compete with inductively coupled plasma mass spectrometry. Various suppliers of energy-dispersive and energy-reflection instruments are listed in Table 1.7. Philips PW1404 energy-dispersive X-ray f luorescence spectrometer The Philips PW1404 is a powerful versatile sequential X-ray spectrometer system developed from the PW1400 series and incorporating many additional hardware and software features that further extend its performance. All system functions are controlled by powerful micro-processor electronics, which make routine analysis a simple, push-button exercise and provide extensive safeguards against operator error. The microprocessor also contains sufficient analytical
Table 1.7 Energy dispersive and total reflection X-ray fluorescence spectrometers
Brief summary of methodologies
31
software to permit stand-alone emergency operation, plus a range of selfdiagnostic service routines. The main characteristics of this instrument are as follows: • • • • •
• • • • • • • • • • • • • •
Identifies all elements from boron to uranium. Choice of side window X-ray tubes allows optimum excitation for all applications. New detectors and crystals bring improved light-element performance. 100kV programmable excitation enhances heavy-element detection. Special calibration features give more accurate results: Auxiliary collimator provides high resolution. Programmable channel mask reduces background. Fast digital scanning speeds data collection. High angular accuracy aids positive identification. Powerful software includes automatic peak labelling. Compact one-cabinet system. Distributed intelligence via five microprocessors. High-frequency generator cuts running costs and improves stability. New high-speed electronics allow operation at 1 million counts per second. System self-selects analytical programs for unknowns. Surface-down sample presentation aids accurate analysis of liquids. Small airlock speeds sample throughput, cuts helium costs. Designed for laboratory automation. Front panel continuously displays system status. New generation software for DEC computers. Computer dialogue in English, French, German and Spanish. Colour graphics simplify results interpretation. Extensive programming, reporting, editing facilities available.
The layout of the Philips PW1404 instrument is shown in Fig. 1.3. Unique among XRF instruments, the EXTRA II TXRF spectrometer yields lower limits of detection in the region of 10pg (1pg = 10-12g) for more than 60 elements e.g. 5–10pg for chlorine, 10–30pg for phosphorus and sulphur and 30–100pg for silicon. All elements upwards from sodium (z = 11) in the periodic table may be determined. The inclusion of twin excitation sources, which may be switched electronically within a few seconds, assures optimum sensitivity for all detectable elements. The applicable concentration range is from per cent to below 1µg L-1. As little as 1µg of sample is sufficient to determine elements at the milligram per litre level; calibration is necessary only once and is carried out during installation. The calibration will remain unchanged for a period of at least 12 months.
Fig. 1.3 Layout of Philips PW1404 energy-dispersive X-ray fluorescence spectrometer.
Brief summary of methodologies
33
Quantitative analysis is simple and uses the method of internal standardization. No external standards are necessary. The method requires no correction of matrix effects for all elements except those in the range sodium to phosphorus. Empirical absorption-enhancement correction models may be applied to these light elements. Sample preparation for solutions and dispersions is very simple, requiring only a micropipette. Complete digestion of materials is not mandatory. Finely divided powders may be analysed providing they are homogeneous. 1.1.9 Isotope dilution methods This technique has found very limited application in water analysis, viz. the determination of chlorocarboxylic acids, cobalamin (vitamin B12) and organically bound nitrogen and phosphorus. 1.1.10 Enzymic assay methods Again there are very few applications, mainly in the area of insecticides/herbicide analysis, viz chlorinated insecticide, polychlorobiphenyl, phosphorus-containing insecticides, carbamate insecticides and herbicides, triazine herbicides, also carbohydrates and phenols. 1.1.11 Chromatographic methods The identification and determination of traces of organic substances in water samples is a subject that has made tremendous advances in recent years. The demands made on water chemists in terms of specificity and sensitivity in carrying out these analyses have become greater and greater with the increasing realization that organic substances from industrial sources are permeating the ecosystem and identification and measurements of minute traces of these are required in potable, river and ground waters and even in rain water . At the same time, measurements in industrial effluent outfalls are necessary in order to control the rate of release of these substances. For the more volatile components of water samples, i.e. those with a boiling point up to about 250°C, gas chromatography has been a favoured technique for several decades. However, with the realization that retention time measurements alone are insufficient to identify organics there has been an increasing move in recent years to connect a gas chromatograph to a mass spectrometer in order to provide unequivocal identifications. Element-specific detectors are another recent development.
34
Introduction
A limitation of gas chromatography is that it cannot handle lessvolatile components of water samples and these comprise a high proportion of the total organics content of the sample. For this reason increasing attention is being paid to the application of high-performance liquid chromatography in water analysis. Again, when positive identifications are required, a mass spectrometer is connected to the outlet of the chromatograph. Supercritical fluid chromatography is a recent development which may find increasing use in water analysis. There are also more specific chromatographic techniques for which instrumentation is now available, such as amino acid analysis, ion-exchange chromatography and ion chromatography. 1.1.11.1 Gas chromatography The numerous substances that have been determined by gas chromatography or gas chromatography–mass spectrometry (marked with asterisks) are listed below: • Hydrocarbons: • Oxygen-containing compounds: • Surface active agents: • Halogen compounds:
• Nitrogen compounds:
• Phosphorus compounds: • Sulphur compounds
• Insecticides:
• Herbicides:
Aliphatic, aromatic, polyaromic. Carboxylic acids, alcohols, glycols, aldehydes, ketones, esters, carbohydrates, phenols*. Anionic, cationic and non-ionic. Saturated and unsaturated* halogen compounds, haloforms*, haloaromics, polychlorobiphenyls,* chlorophenols*, chlorocarboxylic acids, polychloro terphenyls*, chloroanisoles and chloromethyl anisoles*, polychlorodibenzo-p-dioxins and dibenzofurans*, polybromobiphenyls. Aliphatic* and aromatic amines, chloroaromatic amines, aminophenols, nitro compounds*, nitrophenols, nitriles, amides, nitrosamines*, amino acids, chloroamino acids,* ethylenediamine tetraacetic acid, nitriloacetic acid. Alkyl and aryl phosphates. Mercaptans* and disulphides*, dimethyl sulphoxide, t e t r a h y d r o t h i o p h e n , benzthiazole*, 2-mercaptobenzthiazole. Chloro insecticides*, phosphorus insecticides, pyrethrins, carbamate insecticides. Carbamate herbicides, substituted urea
Brief summary of methodologies
• Organometallic compounds of: • Miscellaneous:
• Elements:
35
herbicides, triazine herbicides*, phenoxyacetic acid herbicides, growth regulators. Arsenic, lead, mercury, tin and silicon, Coprostanol, mestranol, ozonizationproducts (ketones), mixtures of organic compounds*, mixtures of pesticides and herbicides*. Total halogen, organic phosphorus.
The basic requirements required of a high-performance gas chromatograph are as follows: •
• •
The sample is introduced to the column in an ideal state, i.e. uncontaminated by septum bleed or previous sample components, without modification due to distillation effects in the needle and quantitatively, i.e. without hold-up or adsorption prior to the column. The instrument parameters that influence the chromatographic separation are precisely controlled. Sample components do not escape detection, i.e. highly sensitive, reproducible detection and subsequent data processing are essential.
There are two types of separation column used in gas chromatography: capillary columns and packed columns. Packed columns are still used extensively, especially in routine analysis. They are essential when sample components have high partition coefficients and/or high concentrations. Capillary columns provide a high number of theoretical plates, hence a very high resolution, but they cannot be used in all applications because there are not many types of chemically bonded capillary columns. Combined use of packed columns of different polarities often provides better separation than with a capillary column. It sometimes happens that a capillary column is used as a supplement in the packed-column gas chromatograph. It is best, therefore, to house the capillary and packed columns in the same column oven and use them routinely and the capillary column is used when more detailed information is required. Conventionally, it is necessary to use a dual-column flow line in packed-column gas chromatography to provide sample and reference gas flows. The recently developed electronic base-line drift compensation system allows a simple column flow line to be used reliably. Recent advances in capillary column technology presume stringent performance levels for the other components of a gas chromatograph as column performance is only as good as that of the rest of the system.
36
Introduction
One of the most important factors in capillary column gas chromatography is that a high repeatability of retention times must be ensured even under adverse ambient conditions. These features combine to provide ±0.01min repeatability for peaks having retention times as long as 2h (other factors being equal). Another important factor for reliable capillary column gas chromatography is the sample injection method. Various types of sample injection ports are available. The split/splitless sample injection port unit series is designed so that the glass insert is easily replaced and the septum is continuously purged during operation. This type of sample injection unit is quite effective for the analysis of samples having high-boiling-point compounds as the major components. In capillary column gas chromatography , it is often required to raise and lower the column temperature very rapidly and to raise the sample injection port temperature. In one design of gas chromatograph, the Shimadzu GC 14-A, the computer-controlled flap operates to bring in the external air to cool the column oven rapidly – only 6min from 500°C to 100°C. This computer-controlled flap also ensures highly stable column temperature when it is set to a nearambient point. The lowest controllable column temperature is about 26°C when the ambient temperature is 20°C. Some suppliers of gas chromatographs are listed in Table 1.8. Instrumentation Shimadzu gas chromatographs This is a typical high-performance gas chromatograph version (Table 1.8 gives further details). The inner chamber of the oven has curved walls for smooth circulation of air; the radiant heat from the sample injection port units and the detector oven is completely isolated. These factors combine to provide demonstrably uniform temperature distribution. (The temperature variance in a column coiled in a diameter of 20cm is less than ±0.75K at a column temperature of 250°C.) When the column temperature is set to a near ambient temperature, external air is brought into the oven via a computercontrolled flap, providing rigid temperature control stability. (The lowest controllable column temperature is 24°C when the ambient temperature is 18°C and the injection port temperature is 250°C. The temperature fluctuation is less than ±0.1K even when the column temperature is set at 50°C.) This instrument features five detectors (Table 1.8). In the flame ionization detector, the high-speed electrometer, which ensures a very low noise level, is best suited to trace analysis and fast analysis using a
Table 1.8 Commercial gas chromatographs
Table 1.8 Continued
Table 1.8 Continued
42
Introduction
capillary column. Samples are never decomposed in the jet, which is made of quartz. Carrier gas, hydrogen, air and make-up gas are separately flowcontrolled. Flow rates are read from the pressure flow-rate curves. In the satellite system, one or more satellite gas chromatographs (GC 14 series) are controlled by a core gas chromatograph. (e.g. GC 16A series). Since the control is made externally, the satellite gas chromatographs are not required to have control functions (the keyboard unit is not necessary). When a GC 16A series gas chromatograph is used as the core, various laboratory-automation-orientated attachments such as a barcode reader and a magnetic-card reader become compatible: a labour-saving system can be built, in which the best operational parameters are automatically set. Each satellite gas chromatograph (GC 14-A series) operates as an independent instrument when a keyboard unit is connected. The IC-card-operated gas chromatography system consists of a GC 14-A series gas chromatograph and a C-R5A Chromatopac data processor. All of the chromatographic and data processing parameters are automatically set simply by inserting the particular IC card. This system is very convenient when one GC system is used for the routine analysis of several different types of samples. One of the popular trends in laboratory automation is to arrange for a personal computer to control the gas chromatograph and to receive data from the GC to be processed as desired. Bilateral communication is made via the RS 232C interface built in a GC 14-A series gas chromatograph. A system can be built to meet requirements. A multidirectional gas chromatography system (multi-stage column system) is effective for analysis of difficult samples and can be built up by connecting several column ovens, i.e. tandem GC systems, each of which has independent control functions such as for temperature programming. The Shimadzu GC 15A and GC 16A systems are designed not only as independent high-performance gas chromatographs but also as core instruments (see above) for multi-gas-chromatography systems (i.e. several gas chromatographs in the laboratory linked to a central management system) or computerized laboratory automation systems. The GC 16A has a keyboard, the GC 15A does not. Other details of these instruments are given in Table 1.8. The Shimadzu GC 8A range of instruments do not have a range of built-in detectors but are ordered either as temperature-programmed instruments with TCD, FID or FPD detectors, or as isothermal instruments with TCD, FID or ECD detectors (Table 1.8).
Brief summary of methodologies
43
Perkin–Elmer Perkin–Elmer supply a range of instruments including the basic models 8410 for packed and capillary work and the 8420 for dedicated capillary work, both supplied on purchase with one of the six different types of detection (Table 1.8). The models 8400 and 8500 are more sophisticated capillary column instruments capable of dual detection operation with the additional features of keyboard operation, screen graphics method storage, host computer links, data handling and compatibility with laboratory automation systems. Perkin–Elmer supply a range of accessories for these instruments including an auto-sampler (AS-8300), an infrared spectrometer interface, an automatic headspace accessory (HS101 and H5-6), an autoinjector device (AI-I) and also a catalytic reactor and a pyroprobe (CDS 190) and automatic thermal desorption system (ATD-550) (both useful for examination of sediments). The Perkin–Elmer 8700, in addition to the features of the models 8400 and 8500, has the ability to perform multi-dimensional gas chromatography. The optimum conditions for capillary chromatography of material heart-cut from a packed column demand a highly sophisticated programming system. The software provided with the model 8700 provides this, allowing methods to be linked so that precolumn and analytical column separations are performed under optimum conditions. Following the first run in which components are transferred from the precolumn to the on-line cold trap, the system will reset to a second method and, on becoming ready, the cold trap is desorbed and the analytical run automatically started. Other applications of the model 8700 system include fore-flushing and back-flushing of the precolumn, either separately or in combination with heart cutting, all carried out with complete automation by the standard instrument software. There are many other suppliers of gas chromatography equipment, some of which are discussed further in Table 1.8. 1.1.11.2 Gas chromatography–mass spectrometry The time has long since passed when one could rely on gas chromatographic or liquid chromatographic data alone to identify unknown compounds in water or other environmental samples. The sheer number of compounds present in such materials would invalidate the use of these techniques, and even in the case of simple mixtures the time required for identification would be too great to provide essential information in the case, for example, of accidental spillage of an organic substance into a water course or
44
Introduction
inlet to a water treatment plant where information is required very rapidly. The practice nowadays is to link a mass spectrometer or ion trap to the outlet of the gas chromatograph or high-performance liquid chromatograph so that a mass spectrum is obtained for each chromatographic peak as it emerges from the separation column. If the peak contains a single substance then computerized library-searching facilities attached to the mass spectrometer will rapidly identify the substance. If the emerging peak contains several substances, then the mass spectrum will indicate this and in many cases will provide information on the substances present. The use of gas chromatography–mass spectrometry grew very rapidly during the early 1970s as discussed by Shackleford and McGuire [32]. The first large-scale application of gas chromatography–mass spectrometry to analysis of environmental pollutants occurred in 1977 when the effluent guidelines division of the EPA, under court order, began collecting data and writing regulations to limit the discharge of pollutants into surface waters. Various workers [33–39] discuss the selection of the EPA priority pollutants and the selection of gas chromatography– mass spectrometry as the technique of choice for their analysis and the problems faced in moving a research technique into production. In addition to the processing work of the EPA, many individual laboratories throughout the world concerned with the analysis of water have set up equipment for gas chromatography–mass spectrometry, and what was once considered to be an expensive instrument, purchased perhaps to solve a single vitally important problem, has become a general workhorse instrument. No water laboratory which aims to be able to solve the kinds of problems thrust upon it can afford to be without a benchtop instrument. Instead, the problem is one of choosing the most appropriate instrumentation for their needs, and this is discussed further below. Instrumentation Finnigan MAT are the main suppliers of this equipment. Finnigan MAT SSQ70 series single-stage quadrupole mass spectrometer This offers premium single-stage performance, with the option of being upgraded to a triple-stage quadrupole system (i.e. the TSQ70). The SSQ70 features a network of distributed microprocessors with more than 1.5Mbyte of memory linked to a powerful DEC 11/73 processor with 2.0Mbyte of memory for data-processing operations. Instrument control links can be displayed in up to eight windows on a colour
Brief summary of methodologies
45
display terminal. The hyperbolic quadrupole analyser gives the SSQ70 a mass range of up to 4000µm; system performance is specified to 200m/z. The cradle vacuum system with three large inlet points at the ion source accommodates a variety of sample inlets such as capillary gas chromatography, thermospray, liquid chromatography, mass spectrometry, supercritical fluid chromatography and solids probe. Standard features of the instrument also include high-performance EI/CI (electron impact/chemical ionization) ion source with exchangeable ion volumes, a PPI NICI with high-voltage conversion dynode multiplier for positive and negative ion detection and fast ion bombardment. Mass spectrometry–mass spectrometr y In high-performance mass spectrometry–mass spectrometry (as opposed to gas chromatography– mass spectrometry) the separator as well as the analysis is performed by the mass spectrometer. One advantage of this technique over combined chromatography–mass spectrometry is that separation is a spatial process rather than being dependent on time. This can lead to improved analysis times and/or greater specificity. Mass spectrometry– mass spectrometry also opens up other areas such as the study of complete structures. This technique has been discussed in detail by Warburton and Millard [40]. Finnigan MAT H-SQ 30 hybrid mass spectrometer–mass spectrometer This instrument combines a reverse-geometry (BE) magnetic section instrument with a quadrupole (QQ) analyser. This hybrid combination provides mass spectrometry–mass spectrometry operation with a high-resolution first stage (BE) and a unit-resolution second stage (QQ). The four available collision regions allow experiments of low (2– 100eV) and high (3keV) collision energy, as well as consecutive collision ion experiments using two separate collision regions. The HSQ 30 is an ideal instrument for structural elucidation studies and ion physics. Finnigan MAT MAT-90 high-mass-high-resolution mass spectrometer This is a veryhigh-performance instrument in which instrument control resides in a multiprocessor system manager leaving only the analytically important parameters to be defined by the operator. It utilizes a completely new concept of ion optics for double focusing and this gives the instrument unmatched performance. The performance of a magnetic sector mass spectrometer depends totally on the ability to focus ions from source to detector. To produce ideal focusing a very wide range of factors must be taken into account. Modern computer simulation techniques have now been extensively applied in this instrument and have resulted in an ion optical design
46
Introduction
closer to the ideal than ever before. This configuration provides for complete image error correction in all planes. System resolution in excess of 50 000 is achieved and excellent performance is obtained at high masses. The instrument features a novel ion source which can be exchanged in a few seconds via vacuum lock. Optimized EI and CI systems are supplied. Optical ionization volumes are available for fast ion bombardment and alternating CC/EI. The MAT-90 analyser has reverse Nier– Johnson geometry allowing metastable studies to be carried out using both first-order and second-order free-field regions in the standard system. To extend the application of metastable techniques, the optical collision cell in the first free-field region can be used. A full range of accessories is available, including direct probe, fast direct probe, thermospray on-line mass spectrometry, automatic direct evaporation, fast ion bombardment, direct chemical ionization and continuous-flow fast ion bombardment. FD/FI device The standard MAT-90 ion source is used for optimized FD/ FI mode by means of the newly designed FD/FI probe. Conversion from electron impact (EI), chemical ionization (CI) or fast ion bombardment (FAB) to FD/FI operation does not require the exchange of the ion source. The FD/FI probe accommodates both the field emitter and the extraction electrodes, mounted at the probe tip. Both are introduced as a unit into the ion source ionization volume exchange lock without breaking vacuum. The fast and simple changeover illustrates the versatility of the MAT-90 with no compromise on the performance. All aspects of system control, data acquisition and processing are carried out in the integrated microprocessor system. The primary processor is a DEC POP 11/73 with 2Mbyte (optional up to 4Mbyte) of main memory with cache and disk cache memory. The user interface is via a high-resolution colour terminal. Standard features also include computer-controlled variable entrance and exit slits, electron multiplier with ±20kV diode, direct coupled capillary gas chromatography–mass spectrometry interface with precise temperature control up to 400°C, data system software including library search, quantification and data handling. Series 700 ion-trap detector The ion-trap detector detects any compound that can be chromatographed; it is a universal detector that can replace several conventional gas chromatography detectors such as the type used in the Varian model 3400 gas chromatogram included in the Finnigan MAT SSA-70 and TSQ-70 instruments. Electron capture, flame ionization, element-specific (etc.) detectors used in the later instruments are not universal in this sense and will not respond to all types of organic compounds, i.e. some compounds
Brief summary of methodologies
47
will be missed. The ion-trap detector obviates the difficulty by responding to all types of organic compounds. In the ion-trap technique one does not have to rely on retention data for identification. The mass spectrum tells you the identity with certainty. Unidentified gas chromatographic peaks are a thing of the past. Complete analysis and identification is done in one run with one detector. This makes the ion-trap detector a very attractive proposition to the water chemist. Various aspects of ion-trap detectors have been discussed by workers at Finnigan MAT and elsewhere [41–63]. During development of the ion-trap detector, it was found that the low voltage previously used for storage encouraged the production and storage of the H O+ and H O+ ions which occasionally led to an 2 3 increase of the M + 1 molecular ions. This problem has been eliminated by adjusting the storage voltage such that the H O+ and 2 H O+ ions are no longer stored. 3 This scanning method produces standard electron impact spectra which can be rapidly searched through the standard National Bureau of Standards (NBS) library (42 222 spectra). In each case the number of ions stored in the trap would be the optimum required to produce a conventional electron impact spectrum. In order that the procedure will not affect quantitative results, ion intensities are stored after application of an adjustment factor which is always related to the true size of the peak as measured by the original fast scan. Scaling is controlled by the computer and the net result is a system with a dynamic range between 104 and 105. The efficiency of the procedure has been evaluated by measuring the signals obtained from injections of a difficult compound in quantities ranging from 10pg to 10ng. Each result has been measured three times and a log/log plot of signals against concentration shown to be a straight line over the entire range with a correlation coefficient >99%. The ion-trap detector may be operated both as a universal detector (whose full scans are stored) or, with the application of multiple-ion monitoring as a specific detector. Because approximately 50% of the ions formed in the trap are analysed, the sensitivity of the instrument in full-scan mode can be much higher than conventional mass spectrometers, in which only 0.1–0.2% of the ions formed may be detected. Thus the instrument can be used to detect 2–5pg (in full scan) of compounds eluting from the column, a performance which compares extremely favourably with those of the most sensitive specific detectors (e.g. the electron capture detector) and easily outstrips that of the flame ionizaton detector. As already indicated, this sensitivity is not achieved at the expense of dynamic range; as the instrument can produce linear calibration graphs for quantities
48
Introduction
within the range 5–10pg to 1000ng on column. This again compares favourably with the performance of the flame ionizaton detector. When operated as a specific detector the ion-trap detector is more sensitive still but not to the extent that would be expected from the performance of other mass spectrometers operated in this mode; in view of the large number of ions monitored in full scan mode there is little more sensitivity to be gained by spending a little extra time scanning a narrow mass range, and the detection limit in this mode is in the region of 1–2pg. The power of the system to overcome the problems associated with co-eluting compounds is demonstrated in conjunction with the use of deuterated (or 13C-labelled compounds) as internal standards. Such techniques could not be used in conventional gas chromatography as the deuterated compounds often co-elute, making quantification difficult if not impossible. With the ion-trap detector, however, it is easily possible to differentiate between the ions arising from the different compounds and the intensities of those ions could then be used for quantification of the compounds involved. The application of such techniques can be shown by the quantitation of anthracene. Chromatograms of ions characteristic of anthracene and its deuterated analogue (m/e 178 and 188 respectively) indicated that the compounds did not in fact elute simultaneously. The retention time of the labelled compound was fractionally less than that of the unlabelled material.
Fig. 1.4 Specific detection. Trace (a) shows a partial chromatogram for Arochlor 1254. Trace (b) shows the mass chromatogram for 292 characteristic of tetrachlorobiphenyls in the mixture. Trace (c) shows the mass chromatogram for 326 characteristic of pentachlorobiphenyls.
Brief summary of methodologies
49
In Fig. 1.4 (a) is shown a partial chromatogram of a complex mixture of chlorinated biphenyls extracted from a water sample. The signals from masses 292 and 326 characteristic of tetra- and pentachlorobiphenyls are shown in Fig. 1.4 (b),(c). The specific detection mode of the ion-trap detector can be used to improve detection limits. This detector can monitor specific masses that are characteristic of compounds of interest. The detector records the signal for only those masses and ignores all others. Interference from other compounds is virtually eliminated with the Finnigan MAT 700 detector – up to 16 different groups of masses can be monitored or a mass range up to 40 masses can be handled. With this flexibility it is possible to monitor only the masses of interest and to improve detection limits. Incos 500 quadrupole mass spectrometer The Incos 500 is a relatively low-cost benchtop instrument as opposed to the research-grade instruments discussed earlier. The gas chromatograph–mass spectrometer transfer lines allow it to be used with either the Hewlett Packard 5890 or the Varian 3400 gas chromatographs. The Incos 500 provides data system control of the gas chromatograph and accessories such as autosampler or liquid sample concentrations. It can be used with capillary wide-bore or packed columns. It performs electron ionization or chemical ionization with positive or negative detection. It also accepts desorption or other solids controls. Finnigan MAT Chem Master Workstation The Chem Master Workstation is a gas chromatography and gas chromatography–mass spectrometry data-processing system that speeds the flow of data through the laboratory and provides essential quality assurance and control review. It is a PC-based integrated hardware–software system that converts gas chromatographic and gas chromatography mass spectrometric data into reliable analytical reports. Model 1020 routine gas chromatograph mass spectrometer This is a cost-effective completely automated system optimized for the routine analysis of complex organic samples. It is specifically designed to meet the needs of an analytical laboratory requiring a gas chromatograph mass spectrometer with the following characteristics:
• • • • • • •
high sample throughput low initial investment low operating costs ease of automation complete software package serviceability field-proven hardware
50
Introduction
The model 1020 software package includes interactive programs specifically designed for complex mixture analysis and advanced automated programs for routine analysis. All system functions are computer controlled with minimal knowledge of mass spectrometry. All gas chromatograph parameters, including temperature program rates and hold times, are controlled by the microprocessor and set through the CRT keyboard. Up to five sets of parameters can be stored on the computer disk for instant recall. The mass spectrometer, when combined with a computer data system, precisely identifies and quantifies each sample component as it elutes from the gas chromatograph. The model 1020 uses an electron ionization source to produce mass fragments, and a quadrupole mass filter, one of the most accurate and cost-effective devices for separating mass fragments. For most applications, excellent spectra are produced with only a few nanograms of sample. When performing single-ion monitoring, picogram and femtogram levels of many compounds can be precisely quantified. In addition to single-ion monitoring, the system’s powerful software permits multiple-ion monitoring of up to 25 mass ranges. This improves sensitivity and reliability of compound identifications by allowing a combination of multiple-ion mass ranges as well as single-ion monitoring to be performed in a single analysis. Mass stability of better than ±0.1µg per day ensures accurate mass assignment. OWA–20/30B organic in water gas chromatograph mass spectrometer This system combines hardware and software features not found in any other low-cost gas chromatography mass spectrometry system. The highly reliable 3000 series electron ionizaton source and quadrupole analyser are used to provide superior mass spectrometer performance. The software is designed with the necessary automation to perform complete quantitative analysis of any target compounds. All routine system operating parameters are adjustable through the computer’s graphics display terminal. The priority interrupt foreground/background operation system allows all data-processing functions to be performed at any time with no limiting effects on data acquisition. Sophisticated dataprocessing programs are readily accessible through a simple commercial structure. The simplicity of the entire system allows complete analysis with minimal operator training. Standard features of this instrument include fully automated gas chromatography mass spectrometry, automated compound analysis and quantification, software, 4–800µ electron impact quadrupole mass spectrometer, high-capacity turbidmolecular pump vacuum system, liquid-sample concentrators for volatile organics in water analysis, a sigma series programmable gas chromatograph, Grob-type split–splitless capillary column injector system, packed column injector with glass jet separator, Nova 4C/53k
Brief summary of methodologies
51
word, 16-bit minicomputer, graphics display terminal, 10Mbyte disk drive, a printer/plotter, an NBS 3100 spectra library, a full-scan or multiple-ion detector and a nine-track tape drive. Options include chemical ionization ion source, direct inlet vacuum lock, programmable solids probe, direct exposure probe, various GC detectors, autosampler, subambient GC operation and a 32Mbyte disk drive. As an example of the application of gas chromatography mass spectrometry, Fig. 1.5 shows a reconstructed ion chromatograph obtained for an industrial waste sample. The Finnigan MAT 1020 instrument was used in this work. Of the 27 compounds searched for, 15 were found. These data were automatically quantified. This portion of the report contains the date and time at which the run was made, the sample description, who submitted the sample to the analyst, followed by the names of the compounds. If no match for a library entry was found, the component was listed ‘not found’. Also shown is the method of quantification and the area of the peak (height could also have been chosen). The large peak at scan 502 (Fig. 1.5) does not interfere with the ability of the software to quantify the sample. Although the compound eluting at scan 502 was not one of the target compounds in the library being reverse-searched, it was possible to identify it by forward-searching the
Fig. 1.5 Reconstructed ion chromatograph of industrial waste sample.
52
Introduction
NBS library present on the system. The greatest similarity was in the comparison of the unknown with the spectrum of benzaldehyde. 1.1.11.3 Headspace gas chromatography For the most volatile substances such as trihalomethanes and halogenated aliphatic compounds, commercial instrumentation is now available for headspace analysis and purge and trap techniques, whereby the gas space above an aqueous sample or the gas purged through an aqueous sample is swept into a gas chromatograph for analysis. Headspace analysis has been applied to the determination of the following types of compounds in water samples: aliphatic hydrocarbons, saturated and unsaturated aliphatic halogen compounds, haloforms and mixtures of volatile organic compounds. Headspace analysis is a method of choice for the determination of volatile compounds in heterogeneous multi-compound water samples. The classic application of this technique is perhaps the determination of trihalomethanes in potable water , but numerous other applications have been found such as the determination of chlorinated aliphatics and volatile hydrocarbons in water. The principle of the analysis is quite simple. The sample is placed in a container leaving a large headspace, which is filled with an inert gas (sometimes under pressure) which also serves as the gas chromatograph carrier gas. Several gas chromatograph manufacturers (Table 1.8) now produce apparatus for headspace analysis. In the past, headspace sampling has often been mentioned in conjunction with low reproducibility. By adding a gas-tight syringe and by employing the latest instrumental refinements to the proven pressure-balanced sampling method, better reproducibility is obtained than that obtained with liquid injection. Typical performance is now better than ±1% relative standard deviation. Under the prevailing equilibrium conditions a proportion of the volatiles in the sample transfer to the gas-filled headspace, which is then withdrawn and analysed by gas chromatography. The HS-100/HS-101 automatic headspace analysers produced by Perkin–Elmer employ a pneumatic pressure-balanced system. The HS100 model is suitable for use with the Perkin–Elmer Sigma 2000 series of gas chromatograph, whilst the HS-101 is designed for use with the 8000 series. With the advanced microprocessor system of the 8000 series and the moving-needle design the HS-101 offers new and important application possibilities unavailable with other headspace injection systems. The HS-101 has a 100-sample storage magazine and is suitable for unattended and night operation. Suppliers of headspace analysers are reviewed in Table 1.9.
Table 1.9 Headspace samplers
54
Introduction
Figure 1.6 shows a gas chromatogram for (a) a mixture of volatile halogenated hydrocarbons and (b) gasoline in water, obtained, respectively, using a packed and a capillary gas chromatographic column. 1.1.11.4 Purge and trap gas chromatography This is an alternative technique to headspace analysis for the identification and determination of volatile organic compounds in water. The sample is purged with an inert gas for a fixed period of time. Volatile compounds are sparged from the sample and collected on a solid sorbent trap – usually activated carbon. The trap is then rapidly heated and the compounds collected and transferred as a plug under a reversed flow of inert gas to an external gas chromatograph. Chromatographic techniques are then used to quantify and identify sample components. Saturated and unsaturated aldehydes, saturated aliphatic halogen compounds and haloforms have been determined in water samples by this technique. OIC Analytical Instrument supply the 4460 A purge and trap concentrator. This is a microprocessor-based instrument with capillary column capability. It is supplied with an autosampler capable of handling 76 sample vials. Two automatic rinses of sample lines and vessel purge are carried out between sample analysis to minimize carryover. Tekmar are another supplier of purge and trap analysis equipment. Their LSC 2000 purge and trap concentrator features glass-lined stainless steel tubing, menu-driven programming with four-method storage and a cryofocusing accessory. Cryofocusing is a technique in which only a short section of the column or a precolumn is cooled. In its simplest form a section of the column near the inlet is immersed in a flask of coolant during desorb. After desorb the coolant is removed and the column allowed to return to the oven temperature. Performance aspects of volatile organics by purge and trap capillary column gas chromatography with flame ionizaton detectors have been discussed by Westendorf [64]. 1.1.11.5 Pyrolysis–gas chromatography Non-volatile organic compounds are not amenable to gas chromatography. However, some types of non-volatile compounds, upon pyrolysis, yield volatile products which are characteristic of the original substance and can be used as the basis of a method for determining these substances. Substances which have been determined using this approach include chlorocarboxylic acids and chlorolignosulphonic acids. The
Brief summary of methodologies
Fig. 1.6 Headspace analysis: (a) halocarbons; (b) gasoline in water.
55
56
Introduction
method is also the basis of one type of procedure for estimating total organic carbon in waters. 1.1.11.6 High-performance liquid chromatography One of the limitations of gas chromatography and consequently of gas chromatography–mass spectrometry is that of all the organic material present in natural water samples, only a small proportion, say as low as 20%, is sufficiently volatile to be separated on gas chromatographic columns operating at even the maximum of their temperature range. Of the 275 compounds in the Appendix III list of the US Environmental Protection Agency, 150 are not amenable to gas chromatographic separation. As a consequence of this there has, in recent years, been a growing interest in applying high-performance liquid chromatography (HPLC) which is not subject to this temperature limitation, to the determination of not only volatile compounds, such as aliphatic and polyaromatic hydrocarbons, saturated and unsaturated aliphatic halogen compounds, haloforms and some esters, phenols and others but also non-volatile components of water (compounds examined by HPLC–mass spectrometry are asterisked: • Hydrocarbons: • Oxygen compounds: • Halogen compounds:
• Nitrogen compounds:
• Sulphur compounds: • Insecticides: • Herbicides:
• Organometallic compounds of: • Miscellaneous:
Aliphatic and polyaromatic types. Carboxyic acids, esters, ethers, phenols, ozonization products (aldehydes). Saturated and unsaturated aliphatic halogen compounds, haloforms, haloaromatic compounds, polychlorobiphenyls, chlorophenols, polychlorodibenzo-p-dioxins and dibenzofurans. Aromatic and chloroaromatic amines, nitro compounds, amides, amino acids, ethylenediamine tetraacetic acid, nitriloacetic acid. Benzthiozole* and 2-mercaptobenzthiozole*, thiafluron. Chloro and phosphorus and carbamate types. Carbamate, and substituted urea and phenoxyacetic acid types, mixtures of pesticides and herbicides*. Lead, tin and copper. Chlorophyll and plant pigments, cobalamin, multiorganic mixtures*.
Brief summary of methodologies
57
Modern high-performance liquid chromatography has been developed to a very high level of performance by the introduction of selective stationary phases of small particle sizes, resulting in efficient columns with large plate numbers per litre. There are several types of chromatographic columns used in highperformance liquid chromatography. Reversed-phase chromatography The most commonly used chromatographic mode in HPLC is reversed-phase chromatography. Reversedphase chromatography is used for the analysis of a wide range of neutral compounds such as carbohydrates, and polar organic compounds. Most common reversed-phase chromatography is performed using bonded silica-based columns, thus inherently limiting the operating pH range to 2.0–7.5. The wide pH range (0–14) of some columns (e.g. Dionex Ion Pac NSI and NS l–5µ columns) removes this limitation, and consequently they are ideally suited for ion-pairing and ionsuppression reversed-phase chromatography, the two techniques which have helped extend reversed-phase chromatography to ionizable compounds. High-sensitivity detection of non-chromophoric organic ions can be achieved by combining the power of suppressed conductivity detection with these columns. Suppressed conductivity is usually a superior approach to using refractive index or low-UV-wavelength detection. Reversed-phase ion-pairing chromatography Typically, reversed-phase ionpairing chromatography is carried out using the same stationary phase as reversed-phase chromatography. A hydrophobic ion of opposite charge to the solute of interest is added to the mobile phase. Samples which are determined by reversed-phase ion-packing chromatography are ionic and thus capable of forming an ion pair with the added counter ion. This form of reversed-phase chromatography can be used for anion and cation separations and for the separation of surfactants and other ionic types of organic molecules. An unfortunate drawback to using silica-based columns is that ion-pairing reagents increase the solubility of silica in water, leading to loss of bead integrity and dramatically reducing column life. Some manufacturers (e.g. Dionex) employ neutral macroporous resins, instead of silica, in an attempt to widen the usable pH range and eliminate the effect of ion-pairing reagents. The technique has been applied to the analysis of phosphoruscontaining insecticides in water. Ion-suppression chromatography Ion suppression is a technique used to suppress the ionization of compounds (such as carboxylic acids) so they will be retained exclusively by the reversed-phase retention
58
Introduction
mechanism and chromatographed as the neutral species. Column packings with an extended pH range are needed for this application as strong acids or alkalis are used to suppress ionization. In addition to carboxylic acids, the ionization of amines can be suppressed by the addition of a base to the mobile phase, thus allowing chromatography of the neutral amine. Ion-exclusion chromatography Haloforms and humic and fulvic acids have been detected by this means. Unlike the pellicular packings used for ion exchange, the packings used in ion exclusion are derived from totally sulphonated polymeric materials. Separation is dependent upon three different mechanisms: Donnan exclusion, steric exclusion and adsorption/partitioning. Donnan exclusion causes strong acids to elute in the void volumes of the column. Weak acids which are partially ionized in the eluent are not subject to Donnan exclusion and can penetrate into the pores of the packing. Separation is accomplished by differences in acid strength, size and hydrophobicity. The major advantage of ion exclusion lies in the ability to handle samples that contain both weak and strong acids. A good example of the power of ion exclusion is the routine determination of organic acids in sea water . Without ion exclusion, the high chloride-ion concentration would present a serious interference. Four basic types of elution system are used in HPLC. This is illustrated below by the systems offered by LKB, Sweden. The isocratic system This consists of a solvent delivery for isocratic reversedphase and gel filtration chromatography. The isocratic system (Fig. 1.7 (a)) provides an economic first step into HPLC techniques. The system is built around a high-performance, dual-piston, pulse-free pump providing precision flow from 0.01 to 5mL min-1. Any of the following detectors can be used with this system: • • • • • • •
fixed-wavelength UV detector (LKB Unicord 2510); variable UV–visible (190–600nm); wavelength monitor (LKB 2151); rapid diode array spectral detector (LKB 2140) (discussed later); refractive index detector (LKB 2142); electrochemical detector (LKB 2143); wavescan EG software (LKB 2146).
Basic gradient system This is a simple upgrade to the isocratic system with the facility for gradient elution techniques and greater functionality (Fig. 1.7 b). The basic system provides for manual operating gradient
Brief summary of methodologies
59
Fig. 1.7 Elution systems supplied by LKB, Sweden (a) isocratic bioseparation system; (b) basic system; (c) advanced chromatography system; (d) inert system.
60
Introduction
techniques such as reversed phase, ion-exchange and hydrophobic interaction chromatography. Any of the detectors listed above under the isocratic system can be used. Advanced gradient system For optimum functionality in automated systems designed primary for reversed-phase chromatography and other gradient techniques, the LKB advanced gradient system is recommended (Fig. 1.7 c). Key features include the following: • • • • •
a configuration that provides the highest possible reproducibility of results; a two-pump system for highly precise and accurate gradient formation for separation of complex samples; separation of complex samples; full system control and advanced method development provided from a liquid chromatography controller; precise and accurate flows ranging from 0.01 to 5mL min-1.
The system is ideal for automatic methods of development and gradient optimization. The inert system By a combination of the use of inert materials (glass, titanium and inert polymers) this system offers totally inert fluidics. Primary features of the system include (Fig. 1.7 d): the ability to perform isocratic or gradient elution by manual means; full system control from a liquid chromatography controller; precise and accurate flows from 0.01 to 5mL min-1. This is the method of choice when corrosive buffers, e.g. those containing chloride or aggressive solvents, are used. Chromatographic detector Details concerning the types of detectors used in HPLC are given in Table 1.10. The most commonly used detectors are those based on spectrophotometry in the region 185–400nm, visible ultraviolet spectrometry in the region 185–900nm, postcolumn derivatization with fluorescence detection (below), conductivity and those based on the relatively new technique of multiple wavelength ultraviolet detectors using a diode array system detector (described below). Other types of detectors available are those based on electrochemical principles, refractive index, differential viscosity and mass detection. Electrochemical detectors These are available from several suppliers (Table 1.10). PSA supply the model 5100A coulochem multi-electrode electrochemical detector. Organics, anions and cations can be detected by electrochemical means.
Table 1.10 Detectors used in HPLC
Table 1.10 Continued
Brief summary of methodologies
63
The Gilson Aspec automatic sample preparation system is a fully automated system for solid-phase extraction on disposable columns and on-line HPLC analysis. The Aspec system offers total automation and total control of the entire sample preparation process including clean-up and concentration. In addition, Aspec can automatically inject prepared samples into on-line HPLC systems. Aspec is designed to receive up to 180 samples. The system is compatible with most standard disposable extraction columns, Analytichem Bond-Elut, Baker SPE, Supelco Supelclean, Alltech Extract Clean, etc. There is a choice of more than 20 different stationary phases. Spectrofluorometric detectors A spectrofluorimeter has been used as a detector in the high-performance liquid chromatographic separation of polyaromatic hydrocarbons in water samples [65–70]. A great improvement in sensitivity and specificity can be obtained by the correct choice of wavelengths. Amino acid analysers This is an example of a dedicated application of HPLC. Measurements of amino acids have been performed in lake water and sea water. Also, measurements of total protein or amino acids are required in river waters and effluents. The most popular current techniques for amino acid analysis rely on liquid chromatography and there are two basic analytical methods. The first is based on ion-exchange chromatography with postcolumn derivatization. The second uses precolumn derivatization followed by reversed-phase HPLC. Derivatization is necessary because amino acids, with very few exceptions, do not absorb in the UV–visible region, nor do they possess natural fluorescence. Each of the major methods has its own particular advantages and disadvantages. Since the variety of available chemistries can be confusing, the method itself should govern the choice that meets requirements, rather than the equipment or systems offered. The optimal method is best selected by a comprehensive and objective review of all commonly used techniques. Overriding criteria which will influence the selection are resolution, sensitivity and speed. Whilst the very best chromatogram for any given method will inevitably be a compromise, only the fullest evaluation of all the alternatives offered will guarantee a correct selection. Certainly, a vast amount of experience has been gained by the widespread use of conventional amino acid analysers. They offer high reliability, accuracy, reproducibility and can separate complex samples. Because conventional analysers can be fully automated, they are widely used in routine analysis. However, the method is limited by the
64
Introduction
sensitivity which can be achieved using ninhydrin as the derivatizing agent. Sensitivity can be increased by using ortho-phthaldialdehyde (OPA) instead, but where extremely high sensitivity is required, HPLC is the method of choice. The other reagents used in HPLC are 9-fluorenyl methoxycarbonyl chloride (FMOC) and phenylisothiocyanate (PITC). Fluorenyl methoxycarbonyl chloride is becoming increasingly popular in protein chemistry research because it reacts with secondary amines and also offers rapid analysis of protein hydrolysates. One aspect governing the choice of method is the sensitivity required. If only a small amount of sample is available, then for the LKB Alpha Plus and for the LKB HPLC instruments, the greatest sensitivity is obtained using the Alpha Plus instrument with the ion-exchange separation and postcolumn derivatization with OPA reagent (108µg) or the HPLC instrument using reversed-phase chromatography and precolumn derivatization with OPA, or FMOC reagents (30–33ng). One leading supplier, LKB, is discussed below. LKB supply two instruments, the LKB 4150 Alpha HPLC, and for analysis requiring higher sensitivity and faster run times the LKB 4151 Alpha Plus. LKB 4150 Alpha This system is a reversed-column chromatograph equipped for precolumn derivatization. The column is made of glass and has solidstate heating. The detection system comprises a dualchannel photometer with a high-temperature reaction coil. A single low-volume longpathlength flow cell is employed. A fluorescence detector is available to provide an approximately 10-fold increase in sensitivity over ninhydrin detection. Refrigerated sample capsule loading is supplied. Powerful programming capability permits the storage of up to 200 methods. Storage facilities for six buffers is supplied. LKB 4151 Alpha Plus Alpha Plus is a fully automated and dedicated analyser. This turnkey system has been carefully designed to give a truly robust chromatography. Stepwise elution with up to five buffers plus flexible temperature control guarantees optimal separations from even the most complex samples. The versatile programmer monitors and controls all instrument functions and a complete fault-detection system assures absolutely safe operation while preserving the integrity of samples. The analysis time for protein hydrolysates is 85min using standard columns. For extra-high resolution a high-resolution lithium cation exchange column is recommended which achieves baseline separation of virtually all 40 amino acids (Fig. 1.8).
Brief summary of methodologies
65
Figure 1.8 Chromatogram of amino acid mixture: sample physiological fluid amino acid standard, 10nmol amino acid in 40 L column – LKB 4418–550 high-resolution column, 270 × 4.6mm packed with lithium from cation-exchange resin; buffers – physiological fluid buffer system (five lithium buffers); flow rate – 20mL h-1 ninhydrin; detection – vis. 570nm and 440nm, 1.0 AUFS.
1.1.11.7 High-performance liquid chromatography–mass spectrometry Applications of this technique are growing and include the analysis of mixtures of pesticides and herbicides, mixtures of various types of organic compounds and mixtures of sulphur compounds (e.g. benzthiazole and 2-mercaptobenzthiazole, alkylbenzene sulphonates [71], polyethylene glycols, nonylphenyl ethoxylates and dioctadecylmethyl ammonium [72, 73]. Hewlett Packard supply the HP 5988A and HP 5987A massselective detectors for use with liquid chromatographs. To date this equipment has been used extensively for identifying and determining non-volatile compounds. The technique has produced further improvements in liquid chromatography mass spectrometry. The particle-beam liquid chromatograph–mass spectrometer uses the same switchable electron impact chemical ionization source and the same software and data system. Adding a gas chromatograph creates a versatile particle–beam liquid chromatograph–gas chromatography–mass spectrometry system that can be switched from liquid chromatography–mass spectrometry to gas chromatography–mass spectrometry in an instant. Based on a new technology, particle-beam enhanced liquid chromatography–mass spectrometry expands a chemist’s ability to analyse a vast variety of substances. Electron impact spectra from the system are reproducible and can be searched against standard or custom libraries for positive compound identification. A simple adjustment to the particle-beam interface is all it takes. The particle-beam system is a simple transport device, very similar to a two-stage jet separator. The solvent vapour is pumped away, while the
66
Introduction
analyte particles are concentrated in a beam and allowed to enter the mass spectrometric source. Here they are vaporized and ionized by electron impact. The different ways a particle-beam liquid chromatograph–mass spectrometer can be configured reflect the versatility of the system in accommodating both the application and the availability of existing instrumentation. The system consists of these elements: • • •
particle-beam interface mounted on the Hewlett Packard 5988A or 5987A mass spectrometer; liquid chromatograph (either the integrated Hewlett Packard 1090 or modular Hewlett Packard 1050); data system (either HP 559970C chem station for a single-instrument operation or the Hewlett Packard 1000 RTE A-series for multiinstrument, multi-tasking, multi-user operation).
This technique is complementary to the thermospray technique. Relative advances of the particle technique over thermospray include library-searchable electron impact spectra, improved reproducibility, easier use and increased predictability over a broad range of compounds. But since a particle beam requires some sample volatility, very large and polar compounds such as proteins may not provide satisfactory results using particle-beam liquid chromatography–mass spectrometry. Additionally, certain classes of compounds such as preformed ions, azo dyes and complex sugars may not yield satisfactory electron impact spectra, but can be run on thermospray. In other words, both liquid chromatography–mass spectrometry techniques complement each other’s limitations and the analyst may want to add both to address a broader range of samples. 1.1.11.8 Ion-exchange chromatography This technique has been used to determine the following types of organic compounds in water: non-ionic surface active agents, saturated aliphatic halogen compounds, organic compounds of arsenic and boron and mixtures of organic compounds. Ion-exchange chromatography is based upon the differential affinity of ions for the stationary phase. The rate of migration of the ion through the column is directly dependent upon the type and concentration of ions that constitute the eluent. Ions with low or moderate affinities for the packing generally prove to be the best eluents. Examples are hydroxide and carbonate eluents for anion separations. The stationary phases commonly used in HPLC are typically derived from silica substrates. The instability of silica outside the pH range 2 to 7.5 represents one of the main reasons why ion-exchange
Brief summary of methodologies
67
separations have not been extensively used in HPLC. To overcome this, some manufacturers (e.g. Dionex) supply columns under the trade name Ion-Pac which contain a packing which is derived from crosslinked polystyrene which is stable throughout the entire pH range. This pH stability allows eluents of extreme pH values to be used so that weak acids such as carbohydrates (and bases) can be ionized. 1.1.11.9 Column coupling capillary isotachoelectrophoresis This technique has found limited applications to the determination of organics in water. These include chlorocarboxylic acids, chlorolignosulphonic acids and growth regulators. The technique offers many similar advantages to ion chromatography, namely multiple-ion analysis, little or no sample pretreatment, speed, sensitivity and automation. Separation capillary columns are made in fluorinated ethylene– propylene copolymer. Detection is achieved by conductivity cells and an a.c. conductivity mode of detection is used for making the separations visible. The driving current is supplied by a unit enabling independent currents to be preselected for the preseparation and final analytical stages. The run of the analyser is controlled by a programmable timing and control unit. The zone lengths from the conductivity detector, evaluated electronically, can be printed on a line printer. To satisfy the requirements for the properties of the leading electrolyte applied in the first stage and, consequently, to decide its composition, two facts have to be taken into account: the pH value of the leading electrolyte needs to be around 4 or less and at the same time the separations of the macroconstituents need to be optimized by means other than adjusting the pH of the leading electrolyte (anions of strong acids). The choice of the leading electrolyte for the second stage, in which the microconstituents were finally separated and quantitatively evaluated, is straightforward, involving a low concentration of the leading constituent (low detection limit) and a low pH of the leading electrolyte (separation according to pK values). 1.1.11.10 Ion chromatography This technique developed by Small et al. in 1975 [74] is usually employed for the separation and determination of mixtures of anions in water. However, some applications have been found for the determination of organic substances in water, viz. mixtures of carboxylic acids, amino acids
68
Introduction
and sulphonates and mixtures of chlorophenols and certain types of organosulphur compounds. The technique uses specialized ion-exchange columns and chemically suppressed conductivity detection. Advances in column and detection technologies have expanded this capability to include wider ranges of anions as well as organic ions. These recent developments, discussed below, provide the chemist with a means of solving many problems that are difficult, if not impossible, using other instrumental methods. Ion chromatography can analyse a wide variety of organic and inorganic anions more easily than either atomic absorption spectrometry or inductively coupled plasma techniques. At the heart of the ion chromatography system is an analytical column containing an ion-exchange resin on which various anions (and/ or cations) are separated before being detected and quantified by various detection techniques, such as spectrophotometry, atomic absorption spectrometry (metals) or conductivity (anions). Ion chromatography is not restricted to the separate analysis of only anions or cations, and, with the proper selection of the eluent and separator columns, the technique can be used for the simultaneous analysis of both anions and cations. The original method for the analysis of mixtures of anions used two columns attached in series packed with ion-exchange resins to separate the ions of interest and suppress the conductance of the eluent, leaving only the species of interest as the major conducting species in the solution. Once the ions were separated and the eluent suppressed, the solution entered a conductivity cell, where the species of interest were detected. The analytical column is used in conjunction with two other columns, a guard column which protects the analytical column from troublesome contaminants, and a preconcentration column. The intended function of the preconcentration column is twofold. First, it concentrates the ions present in the sample, enabling very low levels of contaminants to be detected. Second, it retains non-complexed ions on the resin, while allowing complexed species to pass through. Dionex Series 40001 ion chromatographs Some of the features of this instrument are: • • • •
chromatography module; up to six automated valves made of chemically inert, metal-free material eliminate corrosion and metal contamination; liquid flow path is completely compatible with all HPLC solvents; electronic valve switching, multi-dimensional, coupled chromatography or multi-mode operation;
Brief summary of methodologies
• • • •
69
automated sample clean-up or preconcentration; environmentally isolates up to four separator columns and two suppressors for optimal results; manual or remote control with Dionex Autoion 300 or Autoion 100 automation accessories; individual column temperature control from ambient to 100°C (optional).
Dionex Ion-Pac columns Features are: • •
• • •
Polymer ion-exchange columns are packed with new pellicular resins for anion- or cation-exchange applications; 4µ polymer ion-exchange columns have maximum efficiency and minimum operating pressure for high-performance ion and liquid chromatography applications; New ion-exclusion columns with bifunctional cation-exchange sites offer more selectivity for organic acid separations; Neutral polymer resins have high surface area for reversed-phase ionpair and ion-suppression applications without pH restriction; 5 and 10µ silica columns are optimized for ion-pair, ion-suppression and reversed-phase applications.
Micromembrane suppressor The micromembrane suppressor makes possible detection of non-UV absorbing compounds such as inorganic anions and cations, surfactants, antibiotics, fatty acids and amines in ion-exchange and ion-pair chromatography. Two variants of this exist: the anionic (AMMS) and the cationic (CMMS) suppressor. The micro-membrane suppressor consists of a low dead volume eluent flow path through alternating layers of high-capacity ion-exchange screens and ultra-thin ion-exchange membranes. Ion-exchange sites in each screen provide a site-to-site pathway for eluent ions to transfer to the membrane for maximum chemical suppression. Dionex anion and cation micromembrane suppressors transform eluent ions into less-conducting species without affecting sample ions under analysis. This improves conductivity detection, sensitivity, specificity and baseline stability. It also dramatically increases the dynamic range of the system for inorganic and organic ion chromatography. The high ion-exchange capacity of the MMS permits changes in eluent composition by orders of magnitude, making gradient ion chromatography possible. In addition, because of the increased detection specificity provided by the MMS sample, preparation is dramatically reduced, making it possible to analyse most samples after filtering and dilution.
70
Introduction
Conductivity detector Features include: •
• •
high-sensitivity detection of inorganic anions, amines, surfactants, organic acids, Group I and II metals, oxy-metal ions and metal– cyanide complexes (used in combination with MMS); bipolar-pulsed excitation eliminates the non-linear response with concentration found in analogue detectors; microcomputer-controlled temperature compensation minimizes the baseline drift with changes in room temperature.
UVVis detector Important factors are: •
• • •
high-sensitivity detection of amino acids, metals, silica, chelating agents, and other UV absorbing compounds using either post-column reagent addition or direct detection; non-metallic cell design eliminates corrosion problems; filter-based detection with selectable filters from 214 to 800nm; proprietary dual-wavelength detection for ninhydrin-detectable amino acids and PAR-detectable transition metals.
Optional detectors In addition to the detectors shown, Dionex also offer visible, fluorescence, and pulsed amperometric detectors for use with the series 4000i. Dionex also supply a wide range of alternative instruments, e.g. single channel (2010i) and dual channel (2020i). The latter can be upgraded to an automated system by adding the Autoion 100 or Autoion 300 controller to control two independent ion chromatograph systems. They also supply a 2000i series equipped with conductivity pulsed amperometric, UV-visible and refractometric detectors. 1.1.11.11 Conventional column chromatography Many variants of conventional (non-HPLC) column chromatography have been employed. These lack the sensitivity, resolution and automation of HPLC but are listed below: • Reversed phase adsorption: (Section 1.1.11.6)
• Ion exclusion: (Section 1.1.11.6)
Aliphatic and aromatic hydrocarbons, anionic and non-ionic surface active agents, haloaromatic compounds, polychlorobiphenyls, ethylenediami-ne tetraacetic acid, nitriloacetic acid, phosphorus-containing insecticides. Carboxylic acids, ozonization products (aldehydes).
Brief summary of methodologies
• Ion-pair chromatography: • Partition chromatography: • Size exclusion chromatography: • Electrokinetic chromatography: • Gel permeation chromatography: • Gel filtration chromatography:
71
Mixtures of organic compounds. Carbohydrates. Amides. Esters. Mineral oils, haloforms, humic and fulvic acids. Mixtures of organic compounds.
1.1.11.12 Supercritical fluid chromatography Until recently the chromatographer has had to rely on either gas chromatograpy or HPLC for separations, enduring the limitations of both. Lee Scientific has created a new dimension in chromatography, one which utilizes the unusual properties of supercritical fluids. With the new technology of capillary supercritical fluid chromatography (SFC) the chromatographer benefits from the best of both worlds – the solubility behaviour of liquids and the diffusion and viscosity properties of gases. Consequently, capillary SFC offers unprecedented versatility in obtaining high-resolution separations of difficult compounds. The technique, to date, has found limited but important applications in the determination of organics in waters. These include mixtures of chlorinated insecticides and polychlorobiphenyls, which are notoriously difficult separations to achieve by other forms of chromatography, and also separations of complex mixtures of different types of organic compounds. Beyond its critical point, a substance can no longer be condensed to a liquid, no matter how great the pressure. As pressure increases, however, the fluid density approaches that of a liquid. Because solubility is closely related to density, the solvating strength of the fluid assumes liquid-like characteristics. Its diffusivity and viscosity, however, remain. SFC can use the widest range of detectors available to any chromatographic technique. As a result, capillary SFC has already demonstrated a great potential in applicaton to water, environmental and other areas of analysis. SFC is now one of the fastest-growing analytical techniques. The first paper on the technique was by Klesper et al. [75], but SFC did not catch the analyst’s attention until Novotny et al. [76] published the first paper on capillary SFC. SFC finds its applications in compounds that are either difficult or impossible to analyse by liquid chromatography or gas chromatography. SFC is ideal for analysing either thermally labile or non-volatile nonchromatophoric compounds. The technique will be of interest to water
72
Introduction
chemists as a means of identifying and determining the non-volatile components of water. Most supercritical fluid chromatographs use carbon dioxide as the supercritical eluent, as it has a convenient critical point of 31.3°C and 72.5 atmospheres. Nitrous oxide, ammonia and n-pentane have also been used. This allows easy control of density between 0.2g mL-1 and 0.8g mL -1 and the utilization of almost any detector from liquid chromatography or gas chromatography. Wall [77] has discussed recent developments including timed-split injection, extraction and detection systems in SFC. Timed-split injection Capillary SFC utilizes narrow 50µm or 100µm i.d. columns of between 3 and 20m in length. The internal volume of a 3m × 50µm i.d. column is only 5.8µ1. SFC operates at pressures from 1500lb in 2 to 60001b in -2 (10 335 kPa to 41 340 kPa) and this means that GC injection systems cannot be used. HPLC injection systems are suitable for those pressure ranges, but even using small internalloop injectors the volume introduced to the column is very large compared to the column’s internal volume. To allow injections of about 10–50µl to be introduced to a capillary column, an internalloop LC injector (Valco Inst Switzerland) has been used with a splitter (Fig. 1.9 (a)) which was placed after the valve to ensure that a smaller volume was introduced into the column. This method works well for compounds which are easily soluble in carbon dioxide at low pressures. However, when compounds with higher molecular weights are introduced into the system they are often insufficiently soluble to remain in solution in the depressurization area of the split restrictor. The compounds then reprecipitate in the restrictor and cause a decrease in the internal diameter of the restrictor. Hence this reduces the split ratio and causes more compound to be introduced into the column on the next injection, which means that replicate injections show poor reproducibility. G o o d re p ro d u c i b i l i t y h a s b e e n re p o r t e d f o r c a p i l l a r y supercritical fluid chromatography using a direct injection method without a split restrictor. This method (Fig. 1.9 (b)) utilizes a rapidly rotating internal-loop injector (Valco Inst. Switzerland) which remains in-line with the column for only a short period of time. This then gives a reproducible method of injecting a small fraction of the loop into the column. For this method to be reproducible the valve must be able to switch very rapidly to put a small slug of sample into the column. To attain
Brief summary of methodologies
73
Fig. 1.9 Sample injectors: (a) split valve injector; (b) timed-split and direct valve injector.
this a method called timed-split injection was developed (Lee Scientific). For timed split to operate it is essential that helium is used to switch the valve; air or nitrogen cannot provide sharp enough switching pulses. The injection valve itself must have its internal dead volumes minimized. Dead volumes prior to the valve allow some of the sample to collect prior to the loop, effectively allowing a double slug of sample to be injected which appears at the detector as a very wide solvent peak.
74
Introduction
Detection systems SFC uses detectors from both liquid chromatography and gas chromatography. A summary of detection systems used in SFC has been documented (Later et al. [78]). Electron-capture detector One of the most commonly used detection systems in a gas chromatography laboratory is the electron-capture detector. The first paper (Kennedy and Wall [79]) to be published demonstrating the use of an electron-capture detector with SFC showed that with SFC sensitivity to about 50pg minimum detection limit on column was obtainable. Photoionization detector A paper has been published showing the use of photoionization detectors ([80]). Polyaromatic hydrocarbons are very sensitive using the photoionization detector and the levels detected did not break any new ground in terms of sensitivity. It did inspire NHS Systems (Newton, MA, USA), who market a photoionization detector, to try the detector with a capillary system, interfaced to a Lee Scientific 602 supercritical fluid chromatograph (Lee Scientific, Salt Lake City, Utah, USA). The photoionization detector is, to a certain extent, specific in that only compounds that can be ionized by a UV lamp will give a response. The solvents used were dichloromethane and acetonitrile, both of which should have little response in the photoionization detector. However, a clear sharp solvent peak was observed. The amount detected by this system (0.3pg on column) was below the level which could have been determined using a flame ionization detector. Initial indications show that the photoionization detector may be a very useful detector for people who wish to get lower levels on the supercritical fluid chromatograph and cannot concentrate their sample. Sulphur chemiluminescence detector The flame photometric detector commonly used in gas chromatography for sulphur-specific detection has found little application in SFC. Flame photometric detection is not used in SFC because the initial results obtained showed the response for sulphur-containing species to be very poor. Carbon dioxide has a co-incident emission line which cannot be resolved from the main sulphur line, making flame photometric detection almost useless with SFC and carbon dioxide. Other sulphur detectors do exist, such as the sulphur chemiluminescence detector (CD) (Seivers Research Inc. Colorado, USA). The link to SFC has been investigated. Good sensitivities and chromatograms have been shown for standards and real samples. The detector shows no response to carbon dioxide and
Brief summary of methodologies
75
gives low picogram sensitivities for a wide range of sulphur compounds. The newest developments in SFC instrumentation are the Lee Scientific 602 SFC and 622 SFC/GC. These incorporate the latest advanced technology, the latter being a dual-purpose SFC gas chromatographic instrument. They feature a pulseless high-capacity pump, a high-temperature oven for SFC and gas chromatography, compatibility with packed and capillary columns, high-sensitivity detectors (flame ionization, UV, Fourier transform infrared spectroscopy and mass spectrometry) and newly developed software capable of creating an infinite variety of simultaneous temperature and density or pressure programs. In Fig. 1.10 are shown some SFC chromatograms obtained using the Lee Scientific 501 SFC instrument. In addition to the Lee Scientific, Pierce Chemicals and Severn Analytical in the UK and Valco Instruments in the USA supply equipment for this technique. 1.1.11.13 Thin-layer chromatography This technique has been used extensively for the separation and determination of mixtures of compounds in water by migration on thin layers, usually of silica or alumina. In the case of volatile compounds, such as aliphatic hydrocarbons, care is needed as volatiles may be lost during the separation process. In general, the technique is limited to cases where the identity of the substance to be determined is known, although, in some cases identification of the separated compounds has been achieved by infrared spectrometry or mass spectrometry of elutes of the individual separated spots isolated from the thin-layer plate. Types of compounds which have been determined by thin-layer chromatography are: • Hydrocarbons: • Oxygen compounds: • Surface active agents: • Halogen compounds: • Nitrogen compounds: • Insecticides: • Herbicides:
Aliphatic, aromatic, polyaromatic. Carboxylic acids, aldehydes, lactams, quinones, phenols, phenolic acids. Non-ionics. Saturated aliphatic halogen compounds, polychlorobiphenyls. Nitrophenols, aliphatic amines. Chlorine- and phosphorus-containing insecticides and carbamate insecticides. Carbamate, substituted urea, triazine and phenoxyacetic acid-type herbicides.
76
Introduction
Fig. 1.10 Applications of supercritical fluid chromatography.
• Organometallic compounds of: • Miscellaneous:
Mercury and tin. Chlorophyll and plant pigments, mixtures of various types of organic compounds.
Preconcentration techniques
77
1.2 Preconcentration techniques 1.2.1 Introduction Despite the great strides forward in analytical instrumentation that have been made in the last decade, analysts working in the field of water analysis of non-saline and saline waters find that, frequently, the equipment has insufficient sensitivity to be able to detect the low concentrations of organic substances present in their samples, with the consequence that they have to report less than the detection limit of the method. Consequently, trends upwards or downwards in the levels of background concentrations of these substances in the environment cannot be followed. This is a very unsatisfactory situation which is being made worse by the extremely low detection limits being set in new directives on levels of pollution, issued by the European Community and other international bodies. To overcome the problem, there has been a move in recent years to apply preconcentration to the sample prior to analysis so that, effectively, the detection limit of the method is considerably reduced to the point that actual results can be reported and trends followed. The principle of preconcentration is quite simple. Suppose that we need to determine 5ng L-1 of a substance in a sample and that the best technique has a detection limit of 1µg L-1 (1000ng L-1). To reduce the detection limit to 5ng L-1, we might, for example, pass 1L of the sample down a small column of a substance that absorbs the substance with 100% efficiency. We would then pass down the column 5mL of a solvent or reagent which completely dissolves the substance from the column thereby achieving a preconcentration of 1000/5 = 200. Thus, if the detection limit of the analytical method without preconcentration were 1000ng L -1, then with preconcentration it would be reduced to approximately 5ng L-1. The use of a column is but one of many possible methods of achieving preconcentration. Other methods, listed in Tables 11.11(a)–(e), include preconcentration on macroreticular resins, anion- or cationexchange resins, chelation-solvent extraction techniques, adsorption on solid organic adsorbents or metals or metal oxides, active carbon, organic coprecipitation techniques, solvent extraction, purge and trap and headspace preconcentration. A combination of preconcentration with the newest, most sensitive and, by definition, most expensive analytical techniques now becoming available is achieving previously undreamt-of detection limits at the very time when the requirements for such sensitive analysis are increasing at a rapid pace. Thus the combination of preconcentration with gas chromatography or high performance liquid chromatography
78
Introduction
and, especially, when either of these techniques is coupled with mass spectrometry is enabling exceedingly low concentrations of organics in the microgram per litre range to be determined. Another aspect of preconcentration is, however, worthy of mention, particularly in the case of smaller laboratories which cannot afford to purchase the full range of modern analytical instrumentation. Using older, less sensitive instrumentation, preconcentration will still achieve very useful reductions in detection limits which will be adequate in many but not all instances. Thus, if conventional techniques achieve detection limits of 1 and 5mg L-1, then a 200-fold preconcentration will reduce these limits to approximately 5 and 25µg L-1 and a 1000-fold preconcentration will achieve 1 and 5µg L-1. Most of the types of organic compounds discussed in this book are amenable to preconcentration as is shown in the following list: • Hydrocarbons: • Oxygen compounds:
• Surface active agents: • Halogen compounds:
• Nitrogen compounds: • Phosphorus compounds: • Insecticides: • Herbicides:
• Organometallic compounds of: • Miscellaneous:
Aliphatic, aromatic and polyaromatic and oils. Carboxylic acids, aldehydes, phenols, ethers, ketones, glycol ethers, phthalic esters, carbohydrates, alcohols and dioxans. Anionic and non-ionic types. Saturated aliphatic halogen compounds, haloforms, polychlorobiphenyls, chlorophenols, aromatic halogen compounds. Aromatic amines, nitro compounds, nitroamines, amino acids, quinaldic acid. Alkylphosphates. Chlorine- and phosphorus-containing types, carbamate type and kepone. Carbamate and substituted urea types, chlorophenoxyacetic acid and phenylacetic acid types. Lead, mercury and tin. Humic and fulvic acids, mixtures of pesticides and herbicides, mixtures of different types of organic compounds.
Preconcentration techniques
Table 1.11(a) Adsorption on columns of XAD macroreticular resins
Table 1.11(b) Adsorption on columns of materials other than XAD macroreticular resins
79
80
Introduction
Table 1.11(b) Continued
Table 1.11(c) Coprecipitation techniques
Table 1.11(e) Other techniques
Rationale, analysis of water samples
81
Table 1.11(e) Other techniques
1.2.2 Review of preconcentration techniques Tables 1.11(a) –(e) indicate that by far the most commonly used preconcentration techniques involve the use of columns of XAD, macroreticular resins, Tenax, C -bonded silica, activated carbon, polyolefin 18 films and cation-exchange resins, and, for more volatile components, headspace analysis, purge and trap and gas stripping techniques. 1.2.3 Analytical finishes af ter preconcentration A full review of the preconcentration techniques discussed in the text is given in Table 1.12 in which the technique is cross-referenced with the section number in the text. Details of the type of water sample discussed together with details of the preconcentration techniques and the analytical finish used after preconcentrations are given in this table. 1.3 Rationale, analysis of water samples Tables 1.13(a) –(d) review the methods used for the determination of various types of organic compounds in various types of water samples ranging from natural waters (e.g. rivers, ponds, streams, run-off water, surface water etc.), aqueous precipitation (rain, snow, ice), sea and coastal waters, sewage and trade effluents and miscellaneous waters (e.g. mineral waters and swimming pool waters). The organic compounds discussed cover the whole range of substances that have been known to occur in water samples as a result of industrial, domestic and agricultural inputs; also naturally occurring substances and substances produced as a result of accidental spillages and fires. Particular attention is devoted to the occurrence of agricultural insecticides and herbicides and also of organometallic compounds. Special attention is given to the determination of traces of halogens, nitrogen, phosphorus, sulphur etc. in water samples and to the determination of oxygen demand parameters.
82
Introduction
Table 1.12 Techniques used in the preconcentration of organic compounds in water
Rationale, analysis of water samples Table 1.12 Continued
83
84
Introduction
Table 1.12 Continued
Rationale, analysis of water samples
Table 1.12 Continued
85
86
Introduction
Table 1.12 Continued
Rationale, analysis of water samples
87
Table 1.12 Continued
Note a AAS = atomic absorption spectroscopy; ASV = anodic scanning voltammetry; GLC = gas-liquid chromatography, HPLC = high-performance liquid chromatography, Misc. = miscallaneous; TLC = thin-film liquid chromatography.
In Tables 1.13(a) -(d) analytical techniques are cross-referenced with the organic compound determined, the type of water sample and the section number in the book. If the reader finds that a method is not listed for determining a particular organic compound in the particular type of water sample, then by examination of the table he or she may find a method that is listed under another type of water sample that is applicable to the type of water sample of interest. Thus, if a method is not available for determining a particular organic in say natural water, he or she may find that one is listed under potable water. Obviously this approach will not always be applicable, thus a method listed under trade effluents is not likely to be applicable to the ultra-low-level analysis of potable, natural or sea waters. By far the most extensively used techniques employed in water
Table 1.13(a) Titration, polarographic, voltammetric, spectrophotometric and flow injection methods for the determination of organic compounds in watersa
Table 1.13(a) Continued
Note a Nat = natural waters, pot = potable waters, sew = sewage effluents, trade = trade effluents.
Table 1.13(a) Continued
Table 1.13(b) Atomic absorption, inductively coupled plasma, atomic emission, neutron activation analysis, nuclear magnetic resonance, X-ray fluorescence, isotope dilution and enzymic assay methods for the determination of organics in watersa
Note a Nat = natural waters, pot = potable waters, rain = rain/snow, sew = sewage effluents, trade = trade effluents.
Table 1.13(a) Continued
Table 1.13(c) Gas chromatographic and high-performance liquid chromatographic methods for the determination of organic compounds in watersa
Table 1.13(c) Continued
Table 1.13(c) Continued
Table 1.13(c) Continued
102
Introduction
Table 1.13(d) Liquid chromatographic, superfluid chromatographic and thin-layer chromatographic methods for the determination of organic compounds in watersa
Rationale, analysis of water samples
103
Table 1.13(a) Continued
Note a Nat = natural waters, pot = potable waters, rain = rain/snow, sew = sewage effluents, trade = trade effluents.
analysis are gas chromatography (77 types of organics) and highperformance and conventional liquid chromatography techniques (40 types HPLC, 26 types conventional column chromatography). In a well-equipped laboratory it is mandatory that these techniques be coupled with a mass spectrometric detector in order to achieve a combination of resolution of mixtures, positive identification of separated organics and the high sensitivity that is essential when dealing with environmental water samples. The penetration of mass spectrometers in recent years is indicated by the fact that of the 77 types of organic compounds that have been determined by gas chromatography, in 19 cases mass spectrometric detection is discussed. This trend will, without doubt, continue into the future.
104
Introduction
Another growing technique is supercritical fluid chromatography. Although recent references discuss only polychloroinsecticides, polychlorobiphenyl mixtures and mixtures of other types of organic compounds, there is no doubt that these applications will multiply in the future and that the range of supercritical fluids used (carbon dioxide, and methanol modified carbon dioxide, nitrogen dioxide, ammonia, fluorohydrocarbons) will increase as will the combination of this technique with mass spectrometric identification of separated compounds. For more volatile organic compounds such as aliphatic hydrocarbons, haloforms and saturated and unsaturated low-boiling aliphatic halogen compounds headspace gas chromatography and purge and trap gas chromatography are methods of choice. A technique involving pyrolysis of the organic compound followed by gas chromatography of the pyrolysis products has, to date, found very limited application in the water laboratory (chlorolignosulphonic acids, chlorocarboxylic acids). It is, nevertheless the basis of a well-established method for determining total organic carbon in water. Similarly, the technique of ion chromatography which has extensive application in the determination of anions and cations has very limited application in the determination of organics in water. The substances that have been determined include carboxylic acids, chlorophenols and sulphur-containing organic compounds. Thin-layer chromatography has been applied extensively but is really only of value in preliminary scouting experiments or with types of samples such as sewage and trade effluents where the concentrations of organics present are relatively high, usually in the high milligram per litre range. Regarding routine analysis, which in many cases is amenable to automation, a variety of techniques are available but, as always, in applying these methods the questions of interference effects and sensitivity must be borne in mind. These methods include: 1 2
3
Visible spectrophotometry (32 types of organic compounds), flow injection analysis (six types of organic compounds). Ultraviolet spectroscopy (10 types of organic compounds): unsaturated hydrocarbons, PCBs, phenols, detergents, nitriloacetic acid; humic and fulvic acids, organolead, arsenic and antimony compounds, total organic carbon and dissolved organic carbon. Fluorescence spectrometry (eight types of organic compounds): polyaromatic hydrocarbons, carboxylic acids, phenols, amino acids, carbamate herbicides, humic and fulvic acids, chlorophyll and organoantimony, tin and boron compounds.
References
4
5
105
Polarographic methods (15 types of organic compounds): polyaromatic hydrocarbons, carboxylic acids, aldehydes, esters, cations, PCBs, quinones, detergents, chlorophenols, amides, humic and fulvic acids, phosphorus-containing insecticides, ethylenediamine tetraacetic acid and nitriloacetic acid. Titration methods (13 types of organic compounds): detergents, mercaptans, humic and fulvic acids and chlorine, iodine and organic nitrogen.
Other miscellaneous techniques which have found very limited application in water analysis include the following: 1
2
3
4 5 6 7
8
Infrared spectroscopy (seven types of organic compounds): hydrocarbons, cationic detergents, humic and fulvic acids, mixtures of organics, total organic carbon. Raman spectroscopy (five types of organic compounds): polyaromatic hydrocarbons, phenols, lignosulphonates, phosphoruscontaining insecticides, mixtures of organics. Nuclear magnetic resonance spectrometry (four types of organic compounds): nitrosamines, chlorine-containing insecticides, humic and fulvic acids, mixtures of organic compounds. Neutron activaton analysis (four types of organic compounds): organomercury compounds, chlorine, bromine and iodine. X-ray fluorescence spectroscopy: alkyl and aryl phosphates. Isotope dilution analysis (four types of organic compounds): chlorocarboxylic acids, cobalamin, organic nitrogen and phosphorus. Enzymic assay methods (eight types of organic compounds): carbohydrates, phenols, polychlorobiphenyls, adenosine triphosphate, chlorine and phosphorus-containing insecticides, carbamate herbicides and triazine herbicides. Atomic absorption spectrometry, inductively coupled plasma atomic emission spectrometry and atomic emission spectrometry (13 types of organic compounds): detergents, ethylenediamine tetraacetic acid, nitriloacetic acid, organic compounds of arsenic, lead, mercury, tin, germanium, silica and sulphur.
References 1 2 3 4 5 6
Schalz, L. and Attman, H.J. Z. Analytische Chemie 240 81 (1968) Schwarz, F.P. and Wasik, S.P. Analytical Chemistry 48 524 (1976) Muel, B. and Lacrux, G. Bulletin Chemical Society (London) 2139 (1960) Jager, J. and Fassovitzova, B. Chem. Listy. 62 216 (1968) Khesina, A-Y. and Petrova, J. Spectroscopy U.S.S.R. 18 622 (1973) Monarca, S. Causey, B.S. and Kirkbright, G.F Water Research 13, 503 (1974)
106
Introduction
7 Stepanova, M.I., Il’ina, R.H. and Shaposhnikov, Y.K. Journal of Analytical Chemistry U.S.S.R. 27 1075 (1972) 8 World Health Organization. International Standards for Drinking Water, 3rd edn, Geneva, p. 37(1971) 9 Cathrone, B. and Fielding, M. Proc. Anal. Proceedings Chemical Society (London) 15 155 (1978) 10 Ogan, K., Katz, E. and Slavin, W. Journal of Chromatographic Science 16 517 (1978) 11 Dunn, B.P. and Stich, H.F. Journal of Fisheries Board (Canada) 33 2040 (1976) 12 Kunte, H. Arch. Hyg. Bakt. 151 193 (1967) 13 Ruzicka, J. and Hansen, E.A. Anal. Chim. Acta. 78 145 (1975) 14 Greenfield, S., Jones, I.L. and Berry, C.T. Analyst (London) 89 713 (1964) 15 Wendt, R.H. and Fassel, U.A. Analytical Chemistry 37 920 (1965) 16 Scott, R.H. Analytical Chemistry 46, 75 (1974) 17 Suddendorf, R.F. and Boyer, K.W. Analytical Chemistry 50 1769 (1978) 18 Sharp, B.L. The Conespray Nebulizer, British Technology Group, Patent assignment No. 8 432 338 (1984) 19 Gunn, A.M., Millard, D.L. and Kirkbright, G.F. Analyst (London) 103 1066 (1978) 20 Martusiewicz, H. and Barnes, R.M. Applied Spectroscopy 38 745 (1984) 21 Tikkanen, M.W. and Niemczyk, T.M. Analytical Chemistry 56 1997 (1984) 22 Salin, E.D. and Harlick, G. Analytical Chemistry 51 2284 (1979) 23 Salin, E.D. and Szung, R.L.A. Analytical Chemistry 56 2596 (1984) 24 Thompson, M. and Walsh, J.N. In Handbook of Reductivity Coupled Plasma Spectrometry, Blackie, London and Glasgow, p. 55(1983) 25 Stathan, P.J. Analytical Chemistry 49 2149 (1977) 26 Yaneda, Y. and Horiuchi, T. Dev. Sci. Instr. 42 1069 (1971) 27 Aiginger, H. and Wodbrauschek, P. Nucl. Instruments and Methods 114 157 (1974) 28 Knoth, J., Schwenke, H. and Fresenius, Z. Für Analytisch Chemie 291 200 (1978) 29 Knoth, J., Schwenke, H. and Fresenius, Z. Für Analytisch Chemie 201 7 (1980) 30 Schwenke, H. and Knoth, J. Nuclear Methods 193 239 (1982) 31 Pella, P.A. and Dobbyn, R.C. Analytical Chemistry 60 684 (1988) 32 Shackleford, W.M. and McGuire, J.M. Spectra 10 17 (1986) 33 Telliad, W.A. Spectra 10 4 (1986) 34 Federal Register 44 69466 3 December (1979) 35 Federal Register 45 33066 19 May (1980) 36 Federal Register Method 624 49 43234 26 October (1984) 37 Colby, B.N. Spectra 10 49 (1986) 38 Fish, J.F., Haeberer, A.M. and Kovell, S.P. Spectra 10 22 (1986) 39 Friedman, D. Spectra 10 40 (1986) 40 Warburton, G. and Millard, B. International Labmate xii(7) 87 (1984) 41 Kelly. P.E. Ion trap detection, literature reference list. IDT21. Finnigan, MAT 42 Kelly, P.E. New advances in the operation of the ion trap mass spectrometer. IDT10. Finnigan, MAT 43 Cambell, C. The ion -trap detection for gas chromatography, technology and application. IDT15. Finnigan, MAT 44 Stafford, G.C. Recent improvements in and analytical applications of advanced ion-trap technology. IDT16. Finnigan, MAT 45 Stafford, G.C. Advanced ion -trap technology in an economical detector for GC . IDT20. Finnigan, MAT
References
46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64
65 66 67 68 69 70 71
107
Stafford, G.C. The Finnigan MAT ion-trap mass spectrometer (IT.MS) - new development with ion-trap technology . IDT24. Finnigan, MAT Rordorf, B.F. Comparison of quantitative results for one analysis of 2, 3, 7, 8 TCDD by 4500 Quadrupole MS and 705 ion-trap detector . IDT14. Finnigan, MAT Syka, J.E.P. Positive ion chemical ionization with an ion-trap mass spectrometer . IDT19. Finnigan, MAT Yost, R.A., McClennan, W. and Meuzzelar, H.L.C. Enhanced full scan sensitivity and dynamic range in Finnigan MAT ion-trap detection with new automatic gain control software . IDT22. Finnigan, MAT Camp, C. Ion-trap advancements, higher sensitivity and greater dynamic range with automatic gain control software. IDT23 . Finnigan, MAT Richards, J.M. and Bradford, D.C. Development of a Curie Point Pyrolysis inlet for the Finnigan MAT ion-trap detector. IDT25 . Finnigan, MAT Bishop, P. The ion-trap detector, universal and specific detection in one detector . IDT28. Finnigan, MAT Bishop, P. The use of an ADT50 GLC ion-trap detection combination . IDT36. Finnigan, MAT Bishop, P. Low cost mass spectrometer for GC . IDT42. Finnigan, MAT Campbell, C. and Evans, S. The ion-trap detector – the techniques and its application . IDT29. Finnigan, MAT Olsen, E. Serially interfaced gas chromatography/fourier transform infrared spectrometer/ion-trap spectrometer . IDT35. Finnigan, MAT Allison, J. The hows and whys of ion-trapping . IDT41. Finnigan, MAT Todd, J., Mylechreest, I., Berry, T. and Gaumres, D. Supercritical chromatography mass spectrometry with an ion-trap detector . IDT46. Finnigan, MAT Eichelberg, J.W. and Budd, W.L. Studies in mass spectrometry with an iontrap detector . IDT47. Finnigan, MAT Eichelberg, J.W. and Stivon, L.E. Existence of self chemical ionizaton in the ion-trap detector . IDT48. Finnigan, MAT Genin, E. Le-detecteur a’plegeage d’iou sde chinatogiaphie en phase gaseuse. Technologie et Applications . IDT53. Finnigan, MAT Lebair, M. The use of the IDT51, a low cost GC/MS system for the identification of trace compounds . IDT51. Finnigan, MAT Richards, J.M., McClennan, W.H., Burger, J.A. and Menza, H.H.C. Pyrolysis short column GC/MS using the ITD and ITMS . IDT56. Finnigan, MAT Westendorf, R.G. Tekiman Company P.O. Box. 371856, Cincinnati, OH. 45223-11856, USA. Presented at the 1986 Water Quality Technology Conference of the American Water Works Association, Portland, Oregon, November (1986) Vaughan, C.G., Wheats, B.B. and Whitehouse M.J.J. Journal of Chromatography 28 203 (1973) Lewis, W.M. Water Treatment and Examination 24 243 (1975) Sorrell, R.K. and Reding, R. Journal of Chromatography 185 655 (1975) Sorrell, R.K., Dressman, R. L. and McFerrer, E.F. American Water Works Association Water Quality Technology Conference, Kansas City, MO, December 5 to 7. American Water Works Association, Denver, Colorado , P 3A-3 (1978) Das, B.S. and Thomas, G.H. Analytical Chemistry 50 967 (1978) Schonmann, M. and Kern, H. Varian Instrument Applications 15 6 (1981) Hewlett Packard. Peak, Autumn 10 (1988)
108
72
73 74 75 76 77 78 79 80
Introduction
Rivera, J., Carxash, S., Ventura, F., Fraisse, A. and Des Salees, G. FAB-CADMIKES . Analysis of non ionic surfactants in river and drinking water. Proceedings of 10th International Mass Spectrometry Conference, Swansea , 9–13 September (J.F.F. Todd Editor), John Wiley, Basingstoke (1985) Righton, M.J.G. and Watts, C.D. Water Research Centre , Report ER1194-M. Identification of surfactants in water samples using sublation extraction and fast atom bombardment mass spectrometry . December (1986) Small, H., Stevens, T.S. and Bauman, W.C. Analytical Chemistry 47 1801 (1975) Klesper, E., Corwin, A. and Turner, D. Journal of Organic Chemistry 27 700 (1962) Novotny, M., Springston, P.J. and Lee, M. Analytical Chemistry 53 407A (1981) Wall, R.J. Chromatography and Analysis, John Wiley & Sons, Chichester (1988) Later, D., Bornhof, D., Lee, E., Henion, J. and Wiedholt, R. Liquid Chromatography–Gas Chromatography 1 804 (1987) Kennedy, S. and Wall, R. Liquid Chromatography–Gas Chromatography 2 10 (1988) Sim, P. Elson, C. and Quillaim, M. Journal of Chromatography 445 239 (1988)
Chapter 2
Hydrocarbons
Performing quantitative and qualitative monitoring of low-level volatile hydrocarbons in aqueous solutions is both difficult and essential for environmental and public health operations. These operations include industrial wastewater and process monitoring, site assessment and remediation, and disinfection byproduct control for the drinking water and beverage industries. However, measurements can be problematic in that the analytes of concern are present at low concentrations in a large, potentially interfering background of water. One solution is to separate and concentrate the organic analytes from the water matrix. This has been done with a solvent-free extraction technique. The extraction system uses a silicone-coated silica core fibre mounted on a microsyringe in which the silicone isolates organic analytes that are then thermally injected into a gas chromatograph [401]. Polymeric membranes have also been directly integrated into platforms such as surface-acoustic-wave-based sensors, where frequency shifts are used to detect the presence of chemical analytes in the polymer. These shifts are caused by analyte absorption changing the mass and viscoelastic properties of polymeric materials coated on the quartz crystals [402, 403]. Another application that has been examined extensively in the past decade employs polymer-coated optical wave guides for measurements involving evanescent wave interactions [404– 406]. Inherently weak in nature, the evanescent interactions give information that can be enhanced by using polymeric membranes that separate and concentrate the analytes of concern from their environment. Hydrophobic polymers are frequently employed as cladding material for optical wave guides in both the visible and nearinfrared spectral regions [407–412]. These materials provide mechanical strength environmental protection, and elimination of water interference by excluding water from reaching the optical region of the sensor. Sensors based on the principle known as evanescent film-optic chemical sensors (EFOCS), or fibre-optic evanescent wave sensors (FEWS), hold promise for use in environmental applications.
110
Hydrocarbons
2.1 Aliphatic hydrocarbons 2.1.1 Natural (non-saline) waters 2.1.1.1 Gas chromatograpy Gas chromatography is limited to sufficiently volatile materials. Since petroleum products are readily classified as to type by this technique, the method is highly specific and although substances with similar retention could interfere, the chromatogram profile enables many significant interferences to be noticed and discounted. Gas chromatography has been used to estimate concentrations of volatile petroleum materials in ground water [1]. One-gallon (4.55L) samples of well water were solvent extracted. Iso-octane was employed when a low-boiling petroleum solvent was thought to be the pollutant. A quantitative determination of 20mg L-1 was achieved using n-octane as an internal standard. Direct injection of a petrol-in-water solution, 10mg L-1, was found impractical due to background signal or unknown interference peaks [2]. McAucliffe [3] found various impurity peaks in the direct injection of aqueous solutions of hydrocarbons, which limited sensitivity to about 1mg L-1 of individual hydrocarbons. He employed a 12ft × 1/4 in (3.66m × 6mm) column containing 25% SE–30 on firebrick and a precolumn containing ascarite to absorb the water and improve the sensitivity. Other workers have used similar precolumns to remove water. Two disadvantages of quantitative gas chromatographic analysis of low concentrations are that if different aqueous injection is employed, then background ‘noise’ considerably affects sensitivity, and if a solvent is used for extraction of the oil, impurities in the solvent can be very significant. Headspace analysis and degassing techniques, discussed later, avoid these disadvantages, but are usually applicable only to the more volatile petroleum products. Desbaumes and Imhoff [5] have described a method for the determination of volatile hydrocarbons and their halogenated derivatives in water. Bridie et al. [4] have studied the solvent extraction of hydrocarbons and their oxidative products from oxidized and non-oxidized kerosine– water mixtures, using pentane, chloroform and carbon tetrachloride. Extracts are treated with Florasil to remove non-hydrocarbons before analysis by temperature programmed gas chromatography. From the results reported it is concluded that, although each of the solvents extracts the same amount of hydrocarbons, pentane extracts the smallest amount of non-hydrocarbons. Florasil effectively removes non-
Aliphatic hydrocarbons
111
hydrocarbons from pentane extracts, but also removes 10–25% of aromatic hydrocarbons. However, as the other solvents are less susceptible than pentane to treatment with Florasil, pentane is considered by these workers to be the most suitable solvent for use in determining oil in water. 2.1.1.2 Headspace analysis Khazal et al. [6] and Drodz and Novák [7] examined and compared the methods of headspace-gas and liquid-extraction analysis, comprising the gas chromatography of samples of the gaseous or liquid-extract phases withdrawn from closed equilibrated systems and involving standardaddition quantitation, for the determination of trace amounts of hydrocarbons in water. The liquid-extraction method [8] is more accurate but it yields chromatograms with an interfering background due to the liquid extractant. The sensitivity of determination of volatile hydrocarbons in water is roughly the same for each method, i.e. micrograms per litre. These workers showed that the standard-addition technique is suitable for quantitative determination of trace amounts of hydrocarbons in water, using both headspace analysis and liquid-extraction techniques. Drodz and Novák [9] examined the reliability and reproducibility of qualitative and quantitative headspace analysis of parts per billion of various aliphatic and aromatic hydrocarbons in water using capillary column gas chromatography utilizing a simple all-glass splitless sample injection system. They examined the suitability of the standard-addition method for quantitative headspace gas analysis for concentrations in the condensed phase varying from units to hundreds of parts per billion. With both methods, the chromatographic analysis was carried out at a sensitivity attenuation of 1/16; glass column (180cm × 3mm i.d.) packed with 8.14g of 10% (w/w) Apiezon K on Chromaton N (0.2– 0.25mm); column temperature 80°C; nitrogen carrier gas; flow rated at 26, 29.4 and 200mL min -1 , for nitrogen, hydrogen and air, respectively. The Chromaton N and Apiezon K were products of Lachema and AEI (Manchester, UK) respectively. The headspace method of analysis is less accurate but more sensitive than methods based on liquid extraction. The results in Fig. 2.1 show that with the Drodz 3 [9] method an equilibration time of 10min is adequate for equilibrium between the water sample and the headspace to be achieved. Various other workers [10, 11] have studied the application of headspace analysis to the determination of hydrocarbons in water. McAucliffe [12] determined dissolved individual hydrocarbons in 5mL
112
Hydrocarbons
Fig. 2.1 Chromatograms of 1 mL headspace gas samples, after an equilibration time of (a) 10min, (b) 30min, (c) 2h and (d) 2.5h at 40°C. Peaks; 1, acetone (10ppm in the liquid phase); 2, n-hexane (25ppb); 3, benzene (46ppb); 4, n-heptane (27ppb); 5, toluene (57ppb); 6, n-octane (42ppb); 7, ethylbenzene (79ppb); 8, m-xylene (78ppb); 9, o-xylene (69ppb); 10, n-nonane (44ppb). Column 1, carrier gas N inlet 2 pressure 0.11atm, temperature 70°C. (Reprinted with permission from Drodz and Novák [9]. Copyright (1978) Elsevier Science Publishers BV.)
aqueous samples by injecting up to 5mL of the headspace. For petroleum oils which contain numerous hydrocarbons, very much larger aqueous samples are required. The percentage of hydrocarbons in the gaseous phase, after water containing the hydrocarbons in solution was equilibrated with an equal volume of gas, was found to be 96.7–99.2% for most C –C alkenes. In the case of benzene and 3 8 toluene the values were 18.5 and 21.0%, respectively, indicating that the lower aromatic hydrocarbons may be less amenable to the technique. 2.1.1.3 Gas stripping methods Swinnerton and Linnenbom [13] were the first to examine the applicability of gas stripping methods to the determination of hydrocarbons in water. They determined C –C hydrocarbons by 1 6 stripping them from water with a stream of helium. After gas stripping, the hydrocarbons can either be passed direct to a gas chromatograph or, to increase sensitivity, trapped in a cold trap and then released into the gas chromatograph. Alternatively, the stripped hydrocarbons can be trapped in, for example, active carbon, then released into the gas chromatograph. This method offers the possibility of determining trace amounts of organic compounds in water even below the parts per trillion (ppt) level (1 part in 1012, w/ w), particularly for the most volatile compounds [14]. Many factors,
Aliphatic hydrocarbons
113
such as interference by artefacts because of impurities in the stripping gas, the large amount of water passing the trap, adsorption of less volatile compounds in drying filters, the selection of sorbents and the adsorption and desorption efficiency, are serious drawbacks of the method, particularly for quantitative analysis. However, Grob and co-workers [15–18] reported an impressive improvement of the method by using a closed-loop system, provided with a smallvolume effective charcoal filter, but several precautions are necessary when working at such low concentrations. The complicated procedure and the sophisticated equipment required result in many more-or-less unknown factors and a semiquantitative analysis. In view of the absolute amounts of pollutants involved, their overall results were excellent. Examples of the various types of methods are discussed below. Desbaumes and Imhoff [5] swept volatile hydrocarbons and their halogenated derivatives from water contained in a heated metallic column by a current of purified air and their concentrations were determined with a flame ionizaton detector. The condensed vapours are analysed quantitatively by gas chromatography. Details and diagrams of the equipment are given and the operating procedure is described. Samples must be stored only in glass or stainless steel containers. Substantial losses may still occur if the storage time is less than 10h. Kaiser [20] has described a sensitive degassing technique for trace hydrocarbons in which volatile hydrocarbons up to C (probably 12 higher if the aqueous sample is warmed) are removed from aqueous solution at 20°C by a stream of dry nitrogen during 2–10min, and passed into a gas chromatographic column cooled in liquid nitrogen. After the degassing period was completed, the column temperature was programmed at a rate of 7.5°C min-1, and the hydrocarbons eluted and detected in the usual manner. Substances can be removed from water, other liquids, and solids without any further preparation. The detection limit achieved for individual hydrocarbons in water was 102ppb (10-9wt %). Polak and Lu [19] have described a gas stripping method for the determination of the total amount of volatile but slightly soluble organic materials dissolved in water from oil and oil products. Helium is bubbled through a sample of the aqueous liquid and the gas carries the organic vapours directly to a flame ionization detector. The detector response plotted against time gives an exponential curve, from which the amount of organic material is derived with the aid of an electrical digital integrator. A detector-response factor is required and this is determined with samples prepared by saturating water with hexane or with benzene.
114
Hydrocarbons
Colenutt and Thorburn [22] applied a gas stripping technique to various synthetic and actual samples of hydrocarbons in water. Synthetic solutions of 10µg L -1 n-alkanes from n-octane to nhexadecane prepared by adding acetone solutions of the hydrocarbon to distilled water were put through the procedure. Gas chromatograms were prepared of carbon disulphide extracts. If the solution was analysed almost immediately after preparation a value close to the nominal 10µg L -1 for each component was obtained. However, if the aqueous sample was left exposed in an open laboratory for any length of time, the concentration of the lower molecular weight compounds decreased. Thus the concentrations of the lower alkanes are somewhat suspect in that the evaporation effects prior to sampling are unknown. In Fig. 2.2 is shown a typical gas chromatogram obtained by Colenutt and Thorburn [22] for the extract of a river water sample showing the presence of aliphatic and aromatic hydrocarbons at the 1–10µg L-1 level. These workers found up to 10 times greater concentrations of hydrocarbons in rain water to that found in river water. Drodz et al. [23] used water–air systems with low µg L-1 levels of benzene, toluene, n-decane, n-undecane, and n-dodecane to evaluate the analytical method of repetitive stripping and trapping of analytes. Closed-circuit and open arrangements were investigated to determine the reliability of the method. In a closed circuit, the stripping/ trappingprocess was accomplished under a conservation or an equilibrium regime, whereas in an open arrangement, conservation or pseudoequilibration models of trapping were possible. All the above
Fig. 2.2 Chromatogram of Welsh Harp water sample. Analytical conditions: 10% Carbowax 20M coated on acid-washed silanized Chromosorb W. Nitrogen carrier gas at a flow rate of 30mL min-1. A, n-octane; B, n-nonane; C, benzene; D, n-decane; E, toluene; F, n-undecane; G, ethyl benzene; H, xylenes; I, n-dodecane; J, n-propyl benzene; K, n-tridecane; L, n-tetradecane. (Reprinted with permission from Colenutt and Thorburn [22]. Copyright 1980, Gordon and Breach, London.)
Aliphatic hydrocarbons
115
were used for quantitative analysis. Conservation trapping gave better results when working in an open arrangement. Systematic negative errors of 20 and 40% were obtained for the higher aliphatic hydrocarbons and were attributed to varying matrix effects associated with the adsorption of analytes at the air–water interface. 2.1.1.4 High-performance liquid chromatography Schönmann and Kern [21] have used on-line enrichment for parts per million analysis of aliphatic hydrocarbons and polyaromatic hydrocarbons (PAHs) in water by high-performance liquid chromatography. These workers point out that concentration is a key problem in verifying water quality at parts-per-billion or parts-pertrillion levels. Classical techniques such as freeze drying, extraction and evaporation, steam distillation or preconcentration by adsorption have serious limitations in terms of contamination and sample loss. They used an on-line trace enrichment technique allowing direct high-performance liquid chromatographic analysis of aqueous samples containing very low concentrations of PAHs. This trace enrichment method is based on the affinity of non-polar pollutants for reversed-phase chromatography supports. When aqueous samples are passed through a reversed-phase column these compounds and any other non-polar organic compounds present in the sample are immobilized at the head of the column. When detectable quantities of pollutants have been accumulated on the column, they can be analysed by introducing a mobile phase of the desired eluant strength. The on-column enrichment technique does have some limitations which must be taken into consideration for its successful use. First, compounds which have lower affinity for reversed-phase material may begin to move from the column head during the concentration stage, resulting in peak broadening or analyte breakthrough in cases of large sample volumes or compounds with very low reversed-phase K’ values. This situation limits the utility of on-column concentration for polar phenols, for example. The second limitation of the technique is the time required for introduction of the sample. The pressure limits of most reversed-phase columns require that the sample be introduced at flow rates of a few millilitres per minute, leading to long analysis times for large sample volumes. This problem can be circumvented by concentrating the sample at high flow rate into a low-pressure-drop precolumn, followed by switching in the analytical column to in-line configuration before the introduction of the mobile phase. The oncolumn enrichment method finds its most useful application in the analysis of pollutants in the sub-ppm range in drinking water, ground water, lake and river water.
116
Hydrocarbons
To demonstrate the use of the technique, Schönmann and Kern [21] analysed a lake water sample for PAHs using fluorescence detection with selective filters. The compounds cannot be completely identified by retention times, so an aliquot of the sample was spiked with two compounds suspected to be present. A number of polynuclear aromatics were found in a concentration below 100ng L-1. 2.1.1.5 Infrared spectroscopy Infrared spectroscopy is generally accepted as an excellent technique for determining petroleum products of the order of 1mg or less. Invariably, adsorption intensities of C–H vibrations in aliphatic hydrocarbons are measured, and related to the quantity of the oil present. However, Jeltes [2, 24] points out that when, in very low concentrations, the substances dealt with are in true solution and are aromatic hydrocarbons, infrared spectroscopy is not suitable. In contrast to gas chromatography, the techniques are applicable to nonvolatile petroleum products. The general infrared spectroscopic technique was published as early as 1951 by Simard et al. [25], the oil being extracted from a 1L sample into carbon tetrachloride and the spectra recorded between 3.2 and 3.6µm. Carbon tetrachloride is suitable because it contains no C–H bonds. The intensities of absorptions due mainly to saturated hydrocarbons at 3.38, 3.42 and 3.5µm were determined, and the amount of oil measured down to about 0.1mg L-1 using a standard solution of the oil if possible, or a solution of 37.5% iso-octane, 37.5% cetane and 25% benzene for calibration. Other workers have used this procedure with only slight modifications for the determination of petroleum products in water or soil [26–31]. Similar absorption intensities were measured, i.e. 3.0–3.5µm [27], 3.2–3.6µm [31], 3.3–3.6µm [26], 3.39, 3.42 and 3.51µm [31], 3.25–3.72µm [30]. Various other workers have discussed infrared methods [36–41]. Should any organic matter, such as fatty acids, glycerides, chemical and biochemical oxidation products of petroleum oils, be coextracted into the organic solvent, it can seriously interfere with the determination. To overcome this problem, a prior separation stage has been introduced, involving percolation of the carbon tetrachloride extract through a bed of alumina or Florasil [32–34, 36]. This method is preferred by Hughes et al. [35] who used a modified impeller and a sample bottle immersed in a 30kHz ultrasonic cleaning bath for dispersion of the carbon tetrachloride solvent. Polar materials, for example, carboxylic acids, esters, ketones, phenols and amines, are strongly absorbed, whereas weakly polar hydrocarbons are eluted preferentially and examined by infrared spectrometry.
Aliphatic hydrocarbons
117
Heterocyclic compounds in petroleum products may also be strongly retained. Lindgreen [32] demonstrated that relatively massive contamination by ethanol, acetone, acetic acid, formic acid, oleic acid, vanillin and margarine have little effect on the method, although without prior separation, serious interference occurred. Benyon [34] describes the technique with excellent practical details. A 3.5 to 5L pollution sample was extracted with carbon tetrachloride, and a portion of the extract dried, and then eluted from a Florasil column. The eluant and a blank are examined by infrared analysis and the intensities measured at 3.38µm, and if necessary 3.42 and 3.50µm. A lower limit of 0.1mg kg-1 is quoted with undetermined precision. A variation in the above technique is the formation of an alumina floc in situ, on to which the oil is absorbed [42]. This is filtered off, extracted with carbon tetrachloride, and the spectra produced in the usual maner. Hellmann [43] determined microgram amounts of alkanes in water by extracting 1L of water with as little as 3mL of carbon tetrachloride. This gives a detection limit of l–2µg, with an error of 15–20%. Gruenfeld [44] compared the relative extraction efficiencies of carbon tetrachloride and trichlorotrifluoroethane in the extraction of oils from water and the effects of adding sulphuric acid and sodium chloride were examined by extracting 1L synthetic samples (prepared by emulsifying each of four test oils with tap water) with successive 25mL portions of each solvent. The amount of oil in each extract was determined by measuring the extinction of 2930cm-1. Although the solvents were almost equally effective in extracting the oil, trichlorotrifluoroethane is recommended because of its lower toxicity. Sulphuric acid and sodium chloride improved the extraction efficiency, and complete separation of the oil was achieved with four extractions when 5g of sodium chloride and 5mL of 50% sulphuric aid were added. Mallevialle [45] carried out a systematic study of the factors governing the determination of hydrocarbons in water by extraction with carbon tetrachloride, Florasil chromatography and measurement of infrared adsorption. The method, described below, was unsuitable for aromatic hydrocarbons. Mallevialle [45] standardized on extracting 1L of water, adjusted to pH3 with 10mL of carbon tetrachloride. The water was stirred mechanically for 30min (with a 100-strokes-per-minuite stirring machine mechanism with a backward and forward motion over a 6cm stroke). Efficiency ranged from 65 to 95% for the first extraction and from 95 to 100% for the second; these variations can be explained by the fact that hydrocarbons are in the state of pseudosolutions or microemulsions according to their nature and the presence of surface active elements.
118
Hydrocarbons
Fig. 2.3 Infrared spectrum of crude oil (film on KBr window). (Reprinted with permission from Mallevialle [45]. Copyright (1974) Elsevier Science Ltd, UK.)
Almost all of the hydrocarbons have passed to the organic phase after two extractions from volumes of water of from 1 to 5L. The infrared spectra (KBr disc) of mineral hydrocarbons are of the type illustrated in Fig. 2.3. The peaks of high intensity due to CH, CH 2 and CH in the 2800–3100cm-1 range are characteristic of the aliphatic 3 chains and can therefore be used to measure hydrocarbon content. Measurements were made in infrasil cells with a 10, 20 and 50mm optical path. Mallevialle [45] used a reference mixture as follows: 37.5% trimethylpentane; 37.5% cetane; 25% benzene. The peaks over the range 3000–3100cm-1 are due to the vibrations of the CH groups, particularly of the aromatics; as the intensity of these peaks is much weaker, the aromatics are normally measured by fluorescence or by UV absorption. The peaks at 2962 and 2872 (±10)cm-1 correspond to the CH . In Fig. 2.4 are shown four standard 2 curves: A. B. C. D.
sum of absorptions at 2926 and 2853: CH + CH ; 2as 2s sum of absorptions at 3040, 2962, 2926: CH + CH ; 2as 3as sum of absorptions at 2962 and 2926: CH + CH ; 3as 2as sum of absorptions at 2962, 2926 and 2853: CH + CH 2as
3as
+ CH . 2s
The linearity of the points obtained is satisfactory in each case. Using this method Mallevialle [45] was able to measure as little as 0.1–0.2mg L -1 by means of two extractions with 10mL of carbon tetrachloride from 5L of water (optical path: 10mm) with an accuracy of ±0.05mg L-1. The same result could also be obtained with 1L of water by increasing the optical path to 50mm. The error is considerably reduced ( 370nm; (c) λex 240nm, λem > 470nm. Between 0.5 and 3.0pg of PAHs can be detected by this procedure. 2.3.5.3 Low-temperature spectrofluorimetry using the Shpol’skii ef fect Khesina and Petrova [226] employed this low-level temperature technique in the determination of benzo(a)pyrene and seven other PAHs in extracted waste water, after a preseparation by column chromatography. Other work concerned with the determination of benzo(a)pyrene in environmental waters by this method has been reviewed by Andelmann and Snodgrass [227] and by Andelmann and Suess [228]. 2.3.6 Trade effluents
2.3.6.1 Gas chromatography Searl el al. [229] have described a method for determining PAHs including fluoranthrene, pyrene and benzo(a)pyrene in coke oven effluents by the combined use of gas chromatography and ultraviolet absorption spectrometry. The sample is collected on a silver membrane filter, which is subsequently extracted with cyclohexane in a Soxhlet extractor. After addition of the internal standard, 1,3,5triphenylbenzene, the extract is evaporated to a small volume and subjected to gas chromatography on a column (10ft × 1/8in o.d. (3m × 3mm)) of 2% of SE-30 on acid-washed and dimethylsilylated Chromosorb G (80–100 mesh). The column temperature is programmed from 175 to 275°C at 4°C per minute. The effluent is split so that 15% goes to a flame ionization detector and the rest is trapped in fractions corresponding to the recorded peaks for ultraviolet absorption determination of the compounds in the fractions.
176
Hydrocarbons
Lao et al. [230] described a computerized gas chromatographic-mass spectrometric analysis of PAHs in environmental samples such as coke oven emissions, coal tar, airborne particles, and wood preservative sludges. This method involves a three-step method: preliminary separation of polycyclic aromatic hydrocarbons by solvent and/or column chromatography, identification by gas chromatogram combined with a quadruple mass spectrometer and data processor, and measurement by computerized gas chromatography using internal standards. The efficiencies of different Dexsil gas chromatographic columns were also evaluated during the study. CONCAWE [89] recommended methods described by the Environmental Protection Agency for the determination of PAHs in oil refinery effluents. The method involves extraction of the effluent with methylene dichloride followed by clean-up procedures followed by gas chromatography or liquid chromatography. For the gas chromatographic method no detector is specified; flame ionization seems to be the best choice. Lack of selectivity in the method can lead to interference by compounds that are not completely removed by the clean-up. Higher selectivity can be achieved by use of a photoionization detector. In addition some pairs of the PAH isomers are incompletely separated by the 6in (15cm) column used in this method, while the heavier PAHs often show tailing peaks. Almost all PAH isomers can be separated using a longer column (30in) with the same packing material, or by using a glass capillary column; only benzo(b)fluoranthene–benzo(k)fluoranthene are not resolvable even by the long packed or capillary column approach, both of which are recommended as a final step in preference to a conventional column. The liquid chromatographic procedure [231] uses reversed-phase liquid chromatography with fluorescence detection to separate all 16 PAHs completely. The method is sensitive and so selective as often to allow the method to be applied without clean-up procedure. For gas chromatographic methods, detection limits are about 1µg L-1, whereas for the liquid chromatographic methods, limits are between 1 and 100µg L-1 for two- and three-ring aromatics and below 1µg L-1 for the four-, five- and six-ring compounds. For PAH analysis the extraction with methylene chloride may successfully be substituted by cyclohexane extraction allowing high recoveries by one single extraction. Isolation of PAHs from the water matrix may also conveniently be performed via adsorption using prepacked small high-performance liquid chromatography columns circumventing the problems of emulsification often arising during liquid–liquid extraction. Recently, this technique has been used in a study of polynuclear aromatic hydrocarbons in aqueous effluents from
Polyaromatic hydrocarbons
177
refineries as it was shown to differentiate between certain compounds which were not resolved by gas chromatography–mass spectrometry. 2.3.6.2 High-performance liquid chromatography Kasiske et al. [232] have described a high-performance liquid chromatographic method for six polynuclear aromatic hydrocarbons in trade effluents whose concentration in potable water is regulated by European Union standards, viz. fluoranthene, benzo(e)acetphenanthracene benzo(k)fluoranthene, benzo(def)chrysene, indeno(1,2,3,c,d)pyrene and benzo(ghi)perylene. In this method a 1L sample of water is extracted three times with 30mL of cyclohexane. The combined organic phase is concentrated in a vacuum rotary evaporator to a volume of about 0.5mL and filtered through alumina, activity II. The polycyclic aromatic hydrocarbons adsorbed to the alumina are eluted with 3mL of cyclohexane–benzene (1:1 v/v) and evaporated to dryness. The residue is taken up in 200µL of methanol and 20µL of the latter are injected into the chromatograph. Separation is achieved by elution through a 250 × 4mm i.d. column packed with Nucleosil reversed-phase C18 (particle size 5µm). 2.3.6.3 Miscellaneous Separations on C columns have also been used to analyse PAHs in trade 18 effluents [233, 234]. 2.3.6.4 Preconcentration Kadar et al. [224] studied the efficiencies of Tenax for the removal of PAHs from standard water solutions. The method was applied to waste water samples from an aluminium plant. In this method the water samples were passed through the Tenax column at the rate of about 5mL min-1. Residual water was removed from the Tenax by passing nitrogen gas through the column. The Tenax material was then transferred to a Soxhlet apparatus. PAHs and other organic compounds were extracted by reflux for 4h with 35mL of acetone. The Tenax can then be dried and reused. Total PAHs were then separated from paraffins, naphthenes, acids and phenols on a thinlayer plate prior to determination of the individual PAHs by gas chromatography. Excellent agreement is obtained at the 100ng mL-1 level between the method and a much more time–consuming liquid–liquid extraction technique. The overall recovery of PAHs from water at the 100 and the 10ng mL-1 levels were 90–95.5% and 70–90% respectively. Symons and Crick [414] have conducted a study of the concentration of polynuclear aromatic hydrocarbons from laboratory water and
178
Hydrocarbons
refinery effluents by means of Sep Pak C cartridges. Reversed-phase 18 liquid chromatography with coupled ultraviolet and fluorescence detection is applied to separate and quantify these hydrocarbons. The method was used to determine several polynuclear aromatic hydrocarbons in refinery effluents at the 0.1–50µg L-1 level. 2.3.7 Sewage effluents
2.3.7.1 High-performance liquid chromatography This technique has been applied to the determination of PAHs in sewage amounts down to 2µg L-1 [236, 237]. 2.3.7.2 Miscellaneous Stepanova et al. [235] developed a procedure for the quantitative analysis of a mixture of benzo(a)pyrene and other PAHs in sewage and industrial effluents, based on preliminary thin-layer chromatographic separation and low-temperature spectrophotometric quantification. 2.4. Mineral oil and petroleum spills
2.4.1 Non-saline natural waters
2.4.1.1 Introduction In many areas oil has become the most frequently encountered water pollutant, and oil pollution incidents are becoming more numerous [239, 242, 244, 248]. This reflects the expanding and widespread consumption of petroleum products, a consumption that will continue to increase in the foreseeable future. Oil pollution has harmful effects on aquatic life and lowers the aesthetic appearance of an inland water. Occasionally it necessitates the closure of waterworks intakes. Existing legislation concerning oil pollution cannot be effective unless there are adequate analytical means of detecting oil pollutants, identifying them with regard to their sources and determining their concentration. Many analytical techniques have been proposed, but the references are scattered throughout the literature, and require collation and assessment in relation to inland water pollution. Although marine oil pollution has received much attention in recent years, this has not been the case with oil pollution of inland waters, and
Mineral oil and petroleum spills
179
for some time now there has been need for an assessment of the analytical and related problems in this field. This section discusses these problems. Particular attention is given to the identification of the polluting oil. Wherever oil is produced, stored, transported by vehicle or pipeline, or consumed, there exists a potential source of oil pollution, either directly by surface drains and surface run-off, or indirectly by seepage into the ground. Unlike some pollutants, oil pollution is generally unpredictable as to location and time, and usually exists as a surface phenomenon. Heavy pollution is obviously unwelcome, but even thin ephemeral films representing only small amounts of oil may cause complaints and require investigation if continually present on surface water. The effects of oils on water are manifold, and generally only the acute effects are understood. Long-term chronic effects are slightly comprehended, if at all. Oil, directly or indirectly, seriously lowers the aesthetic appearance of inland waters and interferes with their recreational use. Since oil can readily form a visible thin film of about 7.6 × 10-5mm thickness, a small quantity of oil may be important and produce a heavily polluted appearance. It is often claimed that oil affects the transfer of oxygen into water, but the significance of the effect in the case of thin films is not fully understood. Thick viscous layers do affect the transfer. Photosynthetic activity can be affected by increased reflection and possibly by the absorption of light by the oil. On rivers, where a continuous, prolonged oil slick is improbable, the effect would probably not be significant. Biodegradation of oil in inland waters depletes the oxygen content, but unless frequent oil pollutions occur in a river, such as in heavily industrialized areas, or in non-flowing waters, the depletion would probably do little damage. Most oil products are not considered particularly toxic. Petrol, white spirit and similar volatile products are generally regarded as the most toxic. Heavy surface oil which is emulsified, dissolved, or associated with suspended solids in the body of the water may injure or kill aquatic animal life. Plant life, both aquatic and riparian, can be destroyed by heavy pollution. Prolonged pollution will denude most plant and animal life by a combination of the above-mentioned effects. Long-term toxicity effects are not really understood, although carcinogenicity and general health considerations have been studied in some detail. Recent information suggests that sedimented oils in a lake or river can act as carriers of toxic non-polar organic chemicals, e.g. DDT, owing to the high solvent power of sedimented oils for this type of compound. The significance of this effect in rivers is unknown.
180
Hydrocarbons
Oil contamination is most undesirable in waterworks treatment systems. High concentrations of oil can impair the filters, while even trace quantities can produce taste and odour problems. Practically the total range of petroleum products is encountered as pollutants on inland water. Crude oils are very seldom found inland, but petrol, paraffin, gas oils, heating fuels, lubricating oils, transformer oils, and cutting fluids have given problems. In most areas, heating fuel, due to its widespread industrial and domestic utilization, and diesel fuel are the most commonly occurring oil pollutants. Petrol, although used in greater quantity than most other petroleum products, does not often pollute inland waters. Probably this is due to its high volatility on water surfaces, the strict regulations concerning its storage, and the general public awareness of its dangers. Its relatively high water solubility may be a lesser factor. Lubricating oils give pollution problems, especially in highly industrialized areas. Since lubricating oils are seldom stored or used in large quantities, pollutions tend to be of a smaller nature, but are often responsible for intermittent surface films on inland waters, which are regarded as insignificant. However, the increase in concern over the aesthetic appearance of our inland waters may soon render them significant. Lubricating oils are common pollutants at sewage works. Heavy fuel oils are occasionally met as pollutants, and owing to their viscosity can cause extensive soiling of banks and riparian structures as well as being extremely difficult and troublesome to remove. Natural oils, such as essential oils, vegetable, animal and fish oils, are only infrequently found as significant pollutants. However, when they do occur they can pose extremely difficult problems. The identification procedure for oils in water samples, whether river or ocean can be divided into three stages: 1 2 3
Isolation of the hydrocarbon components from the pollutant sample. Identification of the sample in terms of the petroleum product, for example, crude oil, petroleum, gas oil etc. Identification of the specific source of pollution, such as an individual tanker, tank truck, factory or domestic fuel tank etc.
Stage 2, a general classification of the oil, is often satisfactorily achieved by gas chromatographic techniques possibly coupled with mass spectrometry or infrared spectroscopy applied to a sample of the oil pollutant. Stage 3, the true identification, invariably requires samples from potential sources for comparison with the pollutant. This is often attempted again using gas chromatography, by comparison of the chromatograms, but in a less satisfactory and confident manner. Generally, when the comparisons of chromatograms are reasonably similar, the perpetrator of the pollution accepts liability in the face of
Mineral oil and petroleum spills
181
accumulated circumstantial and scientific evidence and introduces the recommended remedial measures. Existing gas chromatographic techniques can, in the majority of cases, classify petrol, paraffin and light fuel oils. Higher-boiling products with little volatility are not amenable to conventional chromatographic techniques and recourse has to be taken to other techniques such as the use of capillary columns or non-gas–liquid chromatographic techniques. Techniques other than gas chromatography or, more commonly, combinations of techniques have been used to characterize oil spills, as discussed below under separate headings. Nearly every known analytical technique has been used or suggested for oil pollutant identification, but no technique has emerged of such superiority that all the others can be considered redundant. Therefore, earlier attempts of oil characterization have been performed by a multimethod approach; the particular combination of analytical techniques depends on the facilities and the experience existing in a laboratory and the expenditure which is justified to identify any unknown source. Representative examples of the overall approaches are reported in Table 2.13. They include analytical determinations such as the infrared spectra, asphaltene and paraffin contents, etc., that provide a general classification of the pollutants (crude oils, fuel oils, oil sludges, etc.) and others, such a the Ni/V ratio, sulphur content, chromatographic profiles, etc., that permit, by comparison with reference samples, their precise identification. However, another approach involving only one analytical technique, but increasing the number of parameters considered, has been emphasized recently as exemplified in Table 2.14 for trace analysis, infrared spectroscopy and gas chromatography. In these cases a Table 2.13 Overall approaches for identification of oil pollutants
Source: Reprinted with permission from Rasmussen [245]. Copyright (1976) American Chemical Society.
182
Hydrocarbons
Table 2.14 Multiparametic methods for fingerprinting oil pollutants
Source: Reprinted with permission from Rasmussen [245]. Copyright (1976) American Chemical Society.
multiparametric profile is used for identification, instead of a combination of different analytical determinations and pattern recognition techniques have, often, been applied to improve the diagnostic performance. The main requirements that must fulfil these fingerprinting parameters besides their specificity, is that they must remain unaltered during the sea-weathering process affecting the pollutant, namely by evaporation, solution, photo-oxidation and biodegradation. In consequence, both conditions, specificity and stability, need to be investigated in order to evaluate the reliability and the usefulness of any proposed method. General weathering effects occasionally affect identification, and chemists are conscious of the possibility that weathering action on the polluting oil may prevent identification or lead to an incorrect inference. In most cases, since an oil would be unlikely to remain on an inland surface water for any length of time prior to analysis, these effects are probably limited to evaporation of the more volatile components. With practice most analysts can allow for small evaporation effects during the visual inspection of the chromatograms. Little is known of the significance or nature of solution of components on identification. With material of much less volatility than petrol the effect would probably be small. Quantitative analysis of oils polluting the surface of a river is seldom attempted, as the information is not particularly useful. The pollution is obvious, and the value obtained meaningless, because it is impossible quantitatively to determine oil films or slicks, and relate the determination to the total volume of water, the total film area, and the film thickness. Quantitative results for oil are, however, required on effluents entering rivers. The method often used is a gravimetric procedure [83] which determines ‘non-volatile ether-extractable matter’. The method is
Mineral oil and petroleum spills
183
non-specific for petroleum oils, gives erroneous results when soluble oils (cutting oils, etc.) are present, and is inaccurate below the 5mg oil per litre range. Many analysts are dissatisfied with this procedure. Thinlayer chromatography and infrared analysis offer specificity and accuracy in the low-concentration range and will be discussed later. In the case of heavy oil pollution, a sufficient sample of the neat oil is readily taken from the surface. Smaller quantities of oil, for example, thin films and emulsions, can be liquid–liquid extracted with carbon tetrachloride [26, 34], chloroform [246], pentane [1, 34, 246], ether [247], nitrobenzene [2], methanol–benzene (for sediments) [249, 250], hexane [251], toluene–xylene, toluene [252], and iso-octane [253]. Solvents are chosen in order that no significant interference occurs to the components of the pollutant, and that loss of volatile components during evaporation of the solvent, if required, is minimal. For very volatile hydrocarbons in aqueous samples, headspace analysis and degassing techniques are recommended [253, 254]. Goma and Durand used ultrasonically prepared emulsion for their gas chromatographic work, thus avoiding quantitative liquid–liquid extractions [255]. Most authors recommend collection and storage of samples in glass containers. Plastic containers may introduce organic contaminants which may be significant [247], especially with trace amounts of oil. Metal containers may effect subsequent determinations of nickel and vanadium in crude and in residual petroleum oils [252]. Pollutants on the surface can be collected by skimming with a suitable container. Kawahara [246] suggests a glass, wide-mouthed filter funnel fitted with Teflon tubing and a two-way stopcock. The surface oil is ladled into this and is repeatedly separated from the lower aqueous phase. Alternatively, a paint-free dustpan with a stopcock attached to the handle, or even a mop with a fitted wringer, has been employed with success. Samples can be preserved to a limited extent by the addition of solid carbon dioxide to expel air, by refrigeration [246], or by the addition of mercuric chloride which inhibits bacterial activity [254]. 2.4.1.2 Gas chromatography Capillary columns provide greater resolution and therefore more detail for comparison between a polluting oil and suspect sample. The enormous separation power available has been demonstrated in their application to petroleum analysis. Gouw [256] describes a versatile 10m × 0.01in (0.25mm) capillary column coated with CV-101, and its application to the separation of hydrocarbon mixtures in the C –C n-alkane range. 4 58 Columns of 500ft × 0.01in (152m × 0.25mm) coated with 1-octadecene and operated at 30°C have been recommended for identification of crude oils
184
Hydrocarbons
[246]. Hydrocarbons of a selected boiling range, e.g. C –C saturates, were 7 8 collected by prefractionation in a short packed column, trapped, and finally examined on the capillary column. Alkyl-benzenes up to C were 10 examined in a similar manner on an 800ft × 0.01in (240m × 0.25mm) column coated with more polar polyethylene glycol and operated at 60°C. It is claimed that gasoline and other volatile products can be analysed and identified. However, the technique in no way allows for evaporation of volatile components, which would have a serious effect. Columns of the stated length would be impractical for general routine use. A polar liquid phase was found more suitable for studying the major components of petrol, gas oil and diesel oil [2], forming true solutions in water. With such a phase, saturated hydrocarbons tended to elute before aromatic hydrocarbons, which were found to be the principal components in true solution, and therefore their investigation was facilitated, in the case of gas oil and diesel oil, forming true solutions in water. In the case of gas oil and diesel oil, no saturated hydrocarbons could be detected in solution. These authors reached the important conclusion that the determination of the origin of oil components in true aqueous solution could be more difficult because of selective solution of certain components. This effect was likely to apply to the lower distillates, which tended to be relatively more water soluble, rather than the non-volatile petroleum products. Distinction between petrol and gas oil or diesel oil seemed possible, but appeared difficult between similar products such as gas oil and diesel oil. Invariably, dual packed columns have been employed, and one of the earliest articles devoted to the identification of petroleum products is that of Lively [1], who used dual 4ft × 1/4in (1.2m × 6mm) columns packed with 20% SE-30 as the liquid phase an a Chromosorb solid support. Most subsequent workers in this field have employed the same or a similar liquid phase, usually at a lower loading of 4–5% and somewhat different column dimensions, 4–12ft × 1/8–1/4 in (1.2–3.6m × 3–6mm) [257–261]. Chromosorb, or occasionally a similar solid support [259] was usually the solid support utilized after first being acid-washed and treated with hexamethyl disilazine (HMDS) or dimethyl dichlorosilane (DMCS). Liquid phases of similar properties that have been employed are 5% and 10% OV-1, 20% SE-52, 5% E301 [258], and 2.2% [262], 10% [263], and 20% [260] Apiezon L. These liquid phases are essentially non-polar substances, but more polar phases, 5% and 10% polyethylene glycol 1500, have been used for investigations of the watersoluble components in diesel and gas oil and in petrol. In almost all recent publications, flame ionization detectors and temperature programming were employed, the latter especially in the identification of less volatile oils. For routine investigations, isothermal
Mineral oil and petroleum spills
185
conditions have been adopted in order to allow quicker analysis, but at the sacrifice of some fine detail in the chromatogram [260]. Columns containing relatively non-polar liquid phases separate homologous series of hydrocarbons virtually in order of boiling point. In practice, this results in the n-alkanes of petroleum products being separated in order of increasing number of carbon atoms in the molecule. Branched-chain saturated and aromatic hydrocarbons follow a similar pattern, although possibly less precisely. With products such as petrol, paraffin, diesel oil, gas oils and intermediate fuel oils, n-alkanes produce a predominant series of wellseparated peaks on the chromatogram, indicative of the boiling range of the sample. The n-alkanes on the chromatogram usually reside on a mound of poorly resolved branched and cyclic paraffins, aromatic hydrocarbons and heterocyclic compounds. In the case of lubricating, cutting and transformer oils, which are generally lacking in n-alkanes, only fairly featureless mounds are obtained. Pollutants are classified as to type principally by examination of the carbon number range and proportion of the n-alkanes on the chromatogram. Specific n-alkane peaks are usually identified by comparison with standard solutions of some known compounds and a 1% w/v solution has been recommended [34]. Identification of the source of the pollution is achieved by comparison of pollutant and suspect oil chromatograms, usually by matching minor component peaks. It has been stated that while nparaffins tend to be indicative of the processing of a product, the minor components are more specific and can indicate the original crude oil source. With gas chromatography small evaporation effects can be allowed for by expecting irregularities in the volatile component peaks. Detailed salient points involved in classifying and identifying oils have been compiled [258, 264]. Most distillate products, e.g. petrol, paraffin, fuel oils, diesel oil, etc., and even very similar products such as turbojet fuel and kerosene, can be classified in the absence of excessive weathering effects. Various chromatograms have been published: kerosene, turbojet fuel, steam-cracked naphtha; lubricating oil, gas oil, weathered paraffin [264]; white spirit, turpentine substitute, paraffin, 30s fuel oil [260]; standard gasoline; petrol; diesel fuel and gas oil, various gasolines, diesel fuels and aviation fuels [263]. Lubricating, cutting transformer oils etc., heavy fuel oils, asphaltic and bituminous materials are difficult or impossible to classify with any certainty by gas chromatography. A very practical description of the application of gas chromatography to quantitative analysis of petroleum products in aqueous and soil samples is contained in the report by CONCAWE [34]. Well-detailed procedures are given for three hydrocarbon mixtures with the following boiling points:
186
1 2 3
Hydrocarbons
150–450°C, 100–50°C, 0.5mg L-1 the precision is reported as being ±5% for major components and ±10% for others. Jeltes [265] has described a gas chromatographic method for the determination of mineral oil in water in which a true solution of diesel fuel and gas oil in water were analysed by isothermal gas chromatography at 110°C with polyoxyethylene glycol 1550 as stationary phase and Chromosorb W as support. For two-phase oil–water systems, the oil was extracted with carbon tetrachloride and the extracts were analysed by temperature-programmed gas chromatography with SE-30 as stationary phase by a method similar to that of Benyon [34]. Components of mineral oils, i.e. water-soluble types boiling below 300°C, could be identified and determined, and it is possible sometimes to establish the origin of the oil. An interesting gas chromatographic technique of identifying petroleum products, including lubricating oils, is that of Dewitt Johnson and Fuller [261]. A gas chromatograph column effluent was split in order that it could be simultaneously sensed by a double-headed flame photometric detector, specific for both sulphur and phosphorus compounds, and by dual-flame ionization detectors for carbon compounds in general. Since most petroleum products are claimed to contain sulphur and phosphorus compounds, three chromatographic traces were obtained for the products examined. In the cases of lubricating oils, which normally give poorly resolved peaks on chromatograms with flame ionization detection, sulphur and phosphorus detection gave considerably more detail for identifying
Mineral oil and petroleum spills
187
purposes. The apparatus is, however, complex and expensive for routine application. Goma and Durand [255] reported that the sampling errors caused in the injection of water–hydrocarbon mixture on to a gas chromatographic column can be overcome by the addition of acetone for solubilization only when the hydrocarbon chain length is less than 12. Subjection of the sample to ultrasonic vibration (20kHz, 100W) gives a homogenous emulsion which must be used immediately, but it is more satisfactory to add surface active agent (40% of the amount of hydrocarbon if polysorbate 80 is used) and then disperse the sample with ultrasonic vibration. The emulsified sample is injected directly on to a column (4ft × 0.25in (1.2m × 6mm)) of 10% of SE-30 on Chromosorb W HMDS (80– 100mesh) operated at 170–210°C, with hydrogen flame ionization detection. Results are given for the hydrocarbons from C to C . 11 18 Adlard and Matthews [266] applied the flame photometric sulphur detector to pollution identification. A sample of the oil pollutant was submitted to gas chromatography on a stainless steel column (1m × 3mm) packed with 3% of OV-1 on AW-DMCS Chromosorb G (85–100 mesh). Helium was used as carrier gas (35mL min-1) and the column temperature was programmed from 60 to 295°C at 5°C per minute. The column effluent was split between a flame ionization and a flame photometric detector. Adlard and Matthews [266] claim that the origin of oil pollutants can be deduced from the two chromatograms. The method can also be used to measure the degree of weathering of oil samples. Lur’e et al. [267] have described a method which involves extraction of the sample (about 500mL, containing less than 200mg of hydrocarbons) with hexane, concentration of the dried extract to 1mL and gas chromatography of an aliquot (less than 2µL) of this solution. For lower levels of hydrocarbons, 2L of sample is allowed to percolate through a column (13cm × 1cm) on activated carbon, the carbon is dried in air, then the hydrocarbons are extracted with chloroform and the extract is concentrated for gas chromatography. The gas chromatography is carried out on a column (3–4m × 4–5mm) packed with 10% of SE-30 or 20% of 3,3' oxydipropionitrile supported on Chromosorb with helium as carrier and a katharometer detector. Cole [268] found that in the identification of kerosine and aviation fuels, the usual packed columns provided sufficient detail only up to C 13 n-alkane, a range readily altered by evaporation effects. A suitable capillary column was developed which revealed extra detail of minor components above C n-alkane, and consisted of a 45m × 0.25mm 13 stainless steel column coated with OV-101. This gave satisfactory resolution and stability in the 50–310°C temperature range. Naturally abundant saturated terpenoid substances, e.g. farnesane (C ), pristane 15 (C ) and phytane (C ) were determined in fractionated crude oils and 19
20
188
Hydrocarbons
used for identification purposes. The use of a similar column should be applicable to general identification of the commonly occurring oil pollutants of inland waters up to the C –C n-alkane range. This seems 25 30 a logical and valuable extension of the present gas chromatographic techniques. Unfortunately the application to analysis of lubricating oils, etc., does not seem promising. The cost and stability of such a capillary column seems satisfactory for routine uses. Jeltes and Van Tonkelaar [269] compared gas chromatographic and infrared methods for the determination of dissolved mineral oil in water. They saturated various petroleum fractions with water by shaking for 4min, then after 2 days the clear aqueous phases were extracted with carbon tetrachloride. The polar compounds were removed from the carbon tetrachloride extracts by shaking with Florasil and decanting after the Florasil had settled. These extracts were analysed by infrared and gas chromatographic methods, both before and after treatment with Florasil, and also after the two-phase systems had been exposed for 8 weeks to light and air. Gas–liquid chromatography provided the most suitable method for the qualitative and quantitative determination of hydrocarbons; the results were unaffected by Florasil treatment or exposure to light and air. Infrared analysis revealed the presence of dissolved polar compounds. Jeltes and Van Tonkelaar [270] investigated problems of oil pollution, the nature of the contaminants and the chemical methods used for their detection. In particular, the use of gas chromatography to obtain ‘fingerprint’ chromatograms of oil pollutants in water, and of infrared spectrophotometry to determine the oil contents of soils and sediments, is discussed. Kawahara [271] has discussed the characterization and identification of spilled residual fuel oils on surface water using gas chromatography and infrared spectrophotometry. The oily material was collected by surface skimming and extraction with dichloromethane, and the extract was evaporated. Preliminary distinction between samples was made by dissolving portions of the residue in hexane or chloroform. If the residue was soluble in chloroform but not in hexane it was assumed to be crude oil, a grease, a heavy residual fuel oil or an asphalt; if it was soluble in both solvents it was assumed to be very light or heavy naphtha, kerosine, gas oil, white oil, diesel oil, jet fuel, cutting oil, motor oil or cutter stock. The residue was also examined by infrared spectrophotometry; wavenumber values of use for identification purposes are tabulated. Classified, volatile petroleum products such as naphtha, gasoline, jet fuel, kerosine, various fuel oils, crude oil, petroleum jelly and some lubricating oils, were identified by gas chromatography of the residue on, e.g. a column of 5% of DC-200 on Gas Chrom Q temperature programmed from 110 to 224°C at 10° min-1, or a
Mineral oil and petroleum spills
189
capillary column coated with 0.5µm of DC-200 and temperature programmed from 80 to 170°C at 3°C min-1, both operated with helium as carrier gas and flame ionization detection, or a column of 5% of QF-1 and 3% of DC-200 on Gas Chrom Q operated at 180°C with 63Ni electron-capture detection. For heavier products, the pentafluobenzyl derivatives were used for gas chromatography. Lysyj and Newton [272] have described a muticomponent pattern recognition and differentiation method for the analysis of oils in natural waters. The method is based on that described earlier by Lysyj and Newton [273] which depends on the thermal fragmentation of organic molecules followed by gas chromatography. Dried algae and outboardmotor oil were analysed and a specific pattern or numerical ‘fingerprint’ was obtained for each by polygraphic means. The algal pattern comprised three specific peaks and seven peaks common to those of the oil, whereas the oil pattern comprised two specific peaks and seven peaks common to those of the algae. Garra and Muth [274] characterized crude, semi-refined and refined oils by gas chromatography. Separation followed by dual-response detection (flame ionization for hydrocarbons and flame photometric detection for sulphur-containing compounds) was used as a basis for identifying oil samples. By examination of chromatograms, it was shown that refinery oils can be artificially weathered so that the source of oil spills can be determined. Vos et al. [275] have carried out a detailed study of the analysis of oilcontaminated ground water to ascertain the rate of filtration of oil components, and the effects of their biodegradation, under conditions very close to those of natural aquifer. Large-scale lysimeter experiments are reported in a sand dune area where the ground water level could be adjusted with an external overflow device. Details are given of hydrocarbon concentrations determined by adsorption on to Amberlite XAD-4 resins, and investigations using gas chromatography, mass spectroscopy, high-resolution gas chromatography, infrared spectroscopy and ultraviolet spectroscopy. 2.4.1.3 Thin-layer chromatography Thin-layer chromatography has replaced paper chromatography in many applications because of its greater versatility and reproducibility. It has been applied as a rapid method of classifying petroleum and natural and synthetic oils by chromatographing 0.1mg quantities of sample oils [276]. These were spotted on to a silica gel plate and developed with an ascending solvent composed of 70:30 vol/vol chloroform–benzene. After 1h the solvent was evaporated, and the resulting chromatogram evaluated in ultraviolet light (366nm) and the fluorescent areas noted.
190
Hydrocarbons
This was followed by spraying the chromatogram with concentrated sulphuric acid and baking it at 120°C for 15min. Coloured zones formed under visible and ultraviolet light were noted. With this procedure, oils were classified into four groups: 1 2 3 4
hydrocarbon oils (e.g. petroleum products); synthetic ester oils; naturally occurring oils; oils of different composition (e.g. silicone oil and low-molecularweight polyethylene oxides).
The relatively polar solvent, compared to, say, hexane, provides little separation of the hydrocarbon components of petroleum products, and therefore while they can be distinguished from the more polar oils investigated, insufficient information is obtained for differentiation between petroleum products. Krieger [277] employed horizontal silica gel plates and hexane as developing solvent. This resulted in an approximately radial chromatogram which was visualized by spraying with 0.03% fluorescein solution and viewing under ultraviolet light. Crude oil and heavy fuel oil, light petroleum, various tar oils, road tar and olive oil were examined by the technique and some distinction made between them. Natural oil, due to its more polar nature, was not developed, while the remaining products gave generally similar patterns, although in some cases it was possible to match chromatograms of sample and suspect oils. Normal hexane, a relatively non-polar solvent, was adopted by Lambert [278] to produce more separation between petroleum components after ascending development on a silica gel plate. As many as 8–10 samples were chromatographed on a single plate during about 30min, and identification was achieved by comparison of pollutants with standard petroleum materials. Motor oil, light and heavy fuel oil, various greases, and olive oil were among many oils investigated, initially under ultraviolet light and then after spraying with concentrated sulphuric acid–formaldehyde solution. Well-defined coloured zones were obtained with these products with reasonable differentiation between most of the oils examined, although light and heavy fuel oil could be confused. An extension of Lambert’s technique has been achieved [281] in which some commercially available petroleum fractions and greases were examined. Gas oil, motor oil, light fuel oil, motor spirit, paraffin, hydrant grease, submersible pump grease and bitumastic grease were chromatographed on silica gel plates using petroleum ether (b.p. 60–80°C) as developing solvent. Chromatograms were evaluated initially under ultraviolet light
Mineral oil and petroleum spills
191
and characteristic colours and patterns noted. Further detail was collected by spraying the plates with 1% formalin in concentrated sulphuric acid and baking them at 60°C for 16h. Examination under visible and ultraviolet light revealed differently coloured areas. Matthews stated that thin-layer chromatography is highly suited to the rapid identification of the heavier petroleum and coal tar oils, and their residues [279], and has developed a scheme of systematic quantitative analysis [280]. The scheme is based on the varying absorption of oils on different solid supports, for example, silica gel, alumina and kieselguhr, and on the use of solvents of varying polarity, such as petroleum ether (b.p. 40–60°C), ethanol, toluene and chloroform. Ascending thin-layer chromatography was adopted for the scheme, the initial step being separation on alumina and development with acetone. Further thin-layer chromatography over an alternative solid support, or development with a solvent of different polarity follows, depending on the distance travelled by the oil (R value) and its fluorescence under f ultraviolet light (350nm). In principle, material should be classified as to type from R values and its fluorescent patterns, but the author suggests f that in view of small variations encountered in R values standards f should be used. However, the technique is still of value for comparing pollutant oil with a sample from its potential source. Materials were categorized as: 1 2 3 4 5 6
heavy lubricating oils and greases, engineering oils and fuels (a wide range of products from kerosene to light lubricating oil), coal tar products, for example crude tar, anthracene oil, creosotes and pitch, residual and crude petroleum products, other fluorescing oils, for example certain natural oils, non-fluorescing oils, for example liquid paraffin and vegetable oils.
Channel thin-layer chromatography eliminates most of the horizontal spreading of a vertically moving component on the plate, and thereby increases the sensitivity. It has been applied to analysis of oil pollutants [34]. In this modification, two parallel scratches are made 2mm apart on a silica gel coated plate. The sample is applied so that it develops between these scratches with carbon tetrachloride. This permits the thinlayer chromatography of smaller quantities and the resulting rectangular spots are easily measured. The procedure is limited to oils that boil above 150°C. One litre samples are extracted with only 2mL carbon tetrachloride, and loss of any solvent into the water is claimed not to affect the subsequent determination. Therefore exact separation is unnecessary, and the separated extract is chromatographed. On exposure
192
Hydrocarbons
to iodine crystals, brown rectangular areas appear which are easily measured. The developing solvent used, again carbon tetrachloride, probably does not separate saturated and lower aromatic hydrocarbons significantly, and therefore the composition of the oil is less important. A non-volatile gas oil is proposed as a standard hydrocarbon material, and various concentrations are chromatographed to calibrate the plate in terms of area spot (mm 2 ) produced by 0.1mg of the gas oil. Determination of petroleum oils in the range 1–15mg L-1 are possible. Semenov et al. [282] determined small amounts of petroleum products in chloroform extracts of natural water by extracting the sample (200– 500mL) followed by thin-layer chromatography on alumina. He developed the chromatogram with light petroleum–carbon tetrachloride–acetic acid (35:155:1), and examined the plate in UV radiation; the petroleum products exhibit three zones (pale blue, yellow and brown). Each zone is then extracted with chloroform, the fluorescence of the extracts measured and the results referred to a calibration graph. The sensitivity is 0.1mg L -1. The infrared and fluorescence spectra of the zone obtained with various petroleum products are discussed. Sauer and Fitzgerald [283] have described a thin-layer chromatographic technique for the identification of water-borne petroleum oils. Aromatic and polar compounds are removed from the sample by liquid–liquid extraction with acidified methanol, the extract is chromatographed on a silica gel thin-layer plate, and the separated components are detected by their fluorescence under long- and shortwave ultraviolet light. Unsaturated non-fluorescing compounds are detected by iodine staining. The preliminary work of Sauer and Fitzgerald [283] indicated that the resolving power of thin-layer chromatography to separate fluorescing compounds found in oils was poor when the oil, or a dilution thereof, was spotted directly on to the plate; presumably due to the overwhelming complexity of the mixture of compounds generally present in oils. Furthermore, it revealed that only those compounds capable of at least some resistance to chemical alteration due to weathering should be considered. In addition, a simple visualization process such as black light illumination was considered more desirable than more involved techniques such as sulphuric acid charring. For these reasons Sauer and Fitzgerald [283] developed a preliminary fractionation of the starting material: polycyclic aromatic compounds, hereto and polar compounds (N-, O- and S-containing) were isolated from oils and chromatographed. Although these compound types have some solubility in water (and could diffuse into the aqueous layer from an oil spill on water) [101] and are subject to oxidative loss of
Mineral oil and petroleum spills
193
fluorescence, they have provided a satisfactory thin-layer chromatographic fingerprint of oils. Silica gel in conjunction with cyclohexane as solvent [284] was found to be the most suitable absorbent for the separation of saturated from aromatic hydrocarbons in the gas oil to lubricating oil range. Better separation was claimed if the cyclohexane is saturated with less than 0.5% dimethyl sulphoxide. Under the conditions used, paraffins travelled with the solvent front, followed respectively by the monoaromatic, diaromatic, polynuclear aromatic and polar substances, which generally remained on the starting point. Chromatograms can be revealed by ultraviolet light in the normal manner to show aromatic and heterocyclic compounds, and then spraying with dichromate–sulphuric acid reagent to reveal all components, including saturated hydrocarbons [284]. Rhodamine B in fluorescein gives an absorbing spot on a fluorescing background under ultraviolet light at 350 or 254nm. A solution of phosphorus pentachloride in carbon tetrachloride gave some classification of hydrocarbon type in lubricating oils in the following manner [285]: 1 2 3 4
Paraffins–light brown; Monoaromatics–wine red; Diaromatics–dark blue-green; Polar aromatics–dark brown.
Triems [286, 287] classified oils by fractionating the top residue of the oil, freed from compounds boiling at more than 200°C, on silica gel with 2,2,4-trimethylpentane, benzene and acetone as eluants. Three classes of oil were differentiated, viz. paraffinic-naphthenic, naphthenic–aromatic and aromatic–naphthenic. Each class is subdivided into three groups according to its sulphur content. Farrington et al. [288] used column chromatography and thin-layer chromatography to isolate hydrocarbons, arising from marine contamination, from fish lipids. The hydrocarbon extracts were then examined to select those that can be determined by gas chromatography–mass spectrometry, by combinations of spectrophotometric methods, or by wet chemistry. As a screening method gas chromatography was shown to be fairly accurate and precise for hydrocarbons boiling in the range 287–450°C and of suitable polarity. 2.4.1.4 Infrared spectrometry Hellman [289] investigated the possibilities and limits of infrared spectrometry for the determination of mineral and fuel oil in surface waters. The method is applicable to concentrations down to about 0.01–
194
Hydrocarbons
0.2mg L-1. It was used to detect traces of fuel or mineral oil, also to determine the dispersion of oil and its emulsions in water, the biochemical decomposition of mixtures of mineral oil products and the effectiveness of various oil-binding agents and emulsifiers. With oilpolluted samples of soil and ground water, errors were caused by the presence of non-polar hydrocarbons. It was also found that aromatic compounds could not be assessed by infrared spectrophotometry. Geyer et al. [290] tested the feasibility of infrared methods in the case of water containing either toluene, trichloroethylene, or methylene chloride. The characteristic absorption bands of these substances in a carbon tetrachloride extract, following clean-up on an alumina column, were illustrated in the presence or absence of mineral oil. Toluene and oil could only be distinguished with difficulty, while trichloroethylene did not interfere with the quantitative determination of mineral oil; quantitative separation only was possible in the presence of methylene chloride. Peitscher [291] detected and identified traces of oil on surface water. Samples of oil films on surface water were collected with a cloth made of polyester fabric. The cloth was fixed to telescopic rods, so that less accessible sites could be reached and it was kept on the oil interface for periods ranging from several minutes to 1h depending on the amount of oil. The absorbed oil was extracted and analysed by infrared spectrometry. Investigation by techniques including gas chromatography and mass spectrometry facilitated identification of the source of pollution, which could be confirmed by direct comparison of infrared spectra. Differences between spectra for five different types of oil were distinct for a film thickness of 0.2mm but less distinct for a film thickness of 0.1mm. 2.4.1.5 Fluorescence techniques Petroleum products contain many fluorescing components, e.g. aromatic hydrocarbons polycyclic aromatic hydrocarbons and various heterocyclic compounds. The development of improved techniques and instrumentation has led to the use of this technique for the identification of crude and residual oil pollutants in a marine environment [292, 293] and of motor oils and related petroleum products [294–297]. Fluorescence techniques, when they are applicable, are extremely sensitive. The predominant fluorescent substances in petroleum products are polycyclic aromatic hydrocarbons and heterocyclic compounds, and therefore fluorescence should be dependent on the type of oil being examined. This has been found to be an important factor [298], and variations in fluorescence intensities of up to eight times have been found within a small number of lubricating oils examined. Therefore most fluorescent techniques for determining oil in water with useful accuracy demand a sample of the polluting oil, in order that the fluorescence characteristics
Mineral oil and petroleum spills
195
of the oil can be evaluated. Standard hydrocarbons do not show any significant fluorescence. In rivers, lakes and sea water in the vicinity of estuaries, the background fluorescence of naturally occurring organic matter and sewage effluent can be very high, and cause serious interference, while in ground water and in distant sea water it is usually less significant. Nietsch [299–302] and Leoy [303] applied simple fluorescence techniques to the detection of petroleum products in natural waters. Because of the relative purity of the water they were able to detect seepage of the petroleum products into spring water down to about 0.001µg L-1 in favourable circumstances. However, a more realistic level would be about 1mg L -1 with a large undetermined error. The fluorescence of the polluting oil, which could be extracted directly with ether or petroleum ether, or absorbed on to magnesia, was compared to that of solutions of a standard motor oil in water. Only estimates of the oil content are permissible, a limitation that has led to an evaluation of the application of fluorescence to accurate quantitative determination of petroleum oils in water [298]. The authors realized that if any accuracy was to be achieved the oil pollutant must be known to enable its fluorescence characteristics to be ascertained. Aqueous emulsions of various oils were ultrasonically obtained, and their characteristic spectra obtained. The wavelengths of maximum intensity produced by excitation at a suitable wavelength (365nm) were neither characteristic nor reproducible. However, qualitative classification of the oil is claimed as feasible through measurement of three parameters: 1 2 3
the value of total intensity; the width of the curve in nm at half its height; the wavelength corresponding to the centre point of this width.
The qualitative data then allow accurate quantitative calibration of the fluorescence and 1mg L-1 ± 10% is claimed as the lower limit. Unfortunately, practical application to natural waters is complicated, for the authors conclude by reporting that other fluorescing materials present interfere by ‘overshadowing’ the fluorescence of the oil. Also, at lower concentrations, the sensitivity of the method is insufficient. Therefore chemical separation and enrichment procedures are recommended before the method may become applicable. Paper chromatography has been used as a separation technique before application of quantitative fluorescence analysis, to determine bitumens in water [304]. Thrusten and Knight [293] have reported that fluorescence emission spectra obtained by irradiating a cyclohexane solution at 340nm are similar for many crude and semi-refined oils (residual fuels). A
196
Hydrocarbons
maximum emission occurs at about 386nm with slight shoulders at 405 and 440nm. The intensity of the maximum and its shoulders was different for each oil examined, and the ratio of the intensities 440/ 386nm was a further identification parameter. The emission spectra are also dependent on the oil concentration and each oil reacted differently to dilution, which is a third parameter for identification. Evaporation of volatiles in simulated weathering experiments demonstrated that loss of 28–39wt% had little effect. The effect of sunlight and general exposure on a film of oil during 4 days had a very serious effects on the intensities and general shape of many spectra. Immediate sampling is therefore recommended. Freegarde [292] demonstrated that emission spectra from crude oils in solution at 77K and excitation at 250nm contained much more detail. A mixture of fluorescent compounds, such as those in most petroleum materials, gave a different emission spectrum at different excitation (irradiation) wavelengths. An identification technique was developed based on the construction of a contour map of the oil examined, formed by scanning the emission spectrum at 20 excitation wavelengths, and connecting contours of equal fluorescent intensity on a graph of emission wavelength versus excitation wavelength. Synchronous excitation of fluorescence emission is a technique recommended for the rapid identification of motor oils and related petroleum materials [294–296]. Samples of oil in cyclohexane solution are excited at a wavelength which synchronously trailed the plotted emission, usually by about 25nm. The differences between unused motor oils were found to be qualitative rather than quantitative variations, which accords with the findings of Parker and Barnes [297]. Used motor oils varied widely in both aspects, and in this area the technique is promising. Petrols vary distinguishably with octane number and different makes of high-octane petrol tended to vary considerably. As volatility of the product decreases, synchronously excited emission at higher wavelengths predominates. Paraffin, TVO and diesel fuel give spectra seemingly containing less useful detail. Substantial differences existed in the spectra of different types of lubricants, motor oils, gear oils and cutting oil. Differentiations between coal tar pitches may be possible, although bitumens may be less amenable to this technique. The US Environmental Protection Agency [305] has reviewed publications on the identification of petroleum oils in water using fluorescence techniques. John et al. [306] describe how they modified a commercial fluorescence cell to lessen the problems of non-linearity and poor reducibility of spectra when it is used to measure the fluorescence emission from crude oil in water.
Mineral oil and petroleum spills
197
2.4.1.6 Characterization of oils by metals analysis Determination of metals can assist in the characterization of crude oil spillages. Distilled products should not contain these elements. In particular, the ratio of vanadium to nickel in an oil can be characteristic of its source of origin. The ratios of trace elements, particularly V/Ni, have been shown to be unaffected by prolonged weathering effects. Atomic absorption spectrometry Atomic absorption spectrometry has been used to determine vanadium and nickel [307]. Stuart and Branch [307] found that the ratio of vanadium to nickel in oils remains constant during evaporation of lighter fractions, and the value of this ratio as determined by atomic absorption spectrophotometry of the oil diluted with isobutyl methyl ketone can be used to compare oils and to establish their origin. Neutron activation analysis Neutron activation analysis techniques have been reported in several papers dealing with the determination of trace elements in crude oil and petroleum products containing crude oil distillation residues, e.g. marine fuel oil [308, 309]. Bryan et al. [310] developed neutron activation techniques for the identification of metals in oil slicks. Determinations before and after exposure of oils to wave action, elevated temperature, ultraviolet radiation and bacterial attack showed that the concentration of most of the elements was only slightly affected by a minimum exposure, after which they were stable to further exposure. Each trace-element concentration provided an evidence point for comparison of oil samples, and it was found that more than enough evidence points could be determined to distinguish between different oil samples. Determinations of the 34S to 32S ratios also provided a useful evidence point. Lukens et al. [311] discuss procedures in the comparison of slicks with suspected source oils. In particular, neutron activation analysis was used to determine qualitative and quantitative trace-element patterns in a variety of oils, and efforts were made to develop an objective means of comparing these patterns. Nearly 300 trace-element patterns were accumulated, by means of which it can be ascertained, with almost complete certainty, whether or not two samples are of the same oil. Lukens [312] has also published an instruction manual for oil slick identification by trace-element patterns measured by neutron activation analysis. The following steps are described: acquisition of the oil slick sample and of oil samples from suspected sources of the slick;
198
Hydrocarbons
preparation of samples; irradiation of samples and comparator standards under identical conditions; activity measurements on irradiated samples and standards by gamma-ray spectrometry; processing of the spectrometric data in order to identify the trace elements and to determine their concentrations; objective comparison of trace-element patterns of each suspect oil with the slick oil; and estimation of match between suspect and slick oils. Flaherty and Eldridge [313] have discussed the application of neutron activation analysis to the determination of vanadium in oil. The oil sample and standard solution (each 3g) were irradiated for 10min in a neutron flux of about 108 cm-2 s-1, and the gamma-ray spectrum was monitored with an NaI(TI) detector. The maximum deviations of the results over several months were within ±7% for the range 20–2000 ppm of vanadium. Synthetic complexes of manganese, mercury, indium and rhenium with asphaltenes and phthalocyanines, not normally found in petroleum, have been suggested as tracers for crude oils. These were then determined by neutron activation analysis [314]. X-ray f luorescence spectrometry X-ray fluorescence has been employed to determine vanadium and nickel levels in oils [238]. Louis [315] has explained the principles and operation of this technique in the examination of mineral oils and light petroleum fractions. The method is rapid, pretreatment of samples is avoided and improved detection limits are obtained. Kubo et al. [316] have investigated the application of energy dispersive X-ray fluorescence spectrometry to the determination of vanadium, iron, nickel, molybdenum, arsenic, selenium and zinc in fuel oils and shale oils. Gamma-ray spectrometry Gamma-ray spectrometry has been examined [317] and found to compare favourably with conventional chemical techniques for determining metals in oils, but required less time and effort. However, as a computer and an atomic reactor are required the method is instrumentally complicated. Oscillographic polarography Budnikov and Medyantseva [318] determined vanadium and nickel in benzene solutions of bituminous oils by this technique.
Mineral oil and petroleum spills
199
2.4.1.7 Miscellaneous Carsin [319] reviewed sampling equipment used in the determination of hydrocarbon contaminants in natural waters and a new type of sampler in which a floating collector is employed. In this sampler an aspirator pump draws the contaminated water through a bed of oilabsorbent polyurethane foam. Problems of operation of the sampling apparatus, including the behaviour at sea under different weather conditions are discussed. The recovery and subsequent analysis of the oil fraction by infrared spectrometry are also described. Hellmann [320] has shown that certain plastics are permeable to hydrocarbons and may therefore be unsuitable to contain samples of water for analysis. A comparative test by infrared spectrophotometry showed that the hydrocarbon content (initially 1–6mg L -1) of oilcontaminated water fell to about 50% of the initial value after 12h in a plastic container. For very volatile compounds, the loss in the same time may be as much as 90%. Continuous monitoring for oils Although several methods have been developed for continuously monitoring oil or organic materials in water by techniques such as ultraviolet spectroscopy [321] and infrared analysis [322, 323] only a very limited number of published works deal with the detection of surface slicks. This is probably due to the inherent difficulties involved in their design and economic utilization. An oil detector and warning device has been developed [324, 325] mainly for waterworks intakes, but it is also claimed to have applications on inland waters generally. The apparatus automatically detects, reports and records oil films down to 1µm thickness. The whole device floats and therefore tolerates fluctuations in water levels. It operates on flowing waters between 0.1 and 1m s-1 and differentiates between significant and insignificant pollutions. It is claimed to require little maintenance. The detector consists of a rotating plastic disc with fitted electrodes on its periphery. The depth of immersion is adjustable. Oil is removed from the water during rotation of the disc, which causes a decrease in conductance across the electrodes, and finally by a cleaning device before reimmersion. Reflection measurements were used by Goolsby [326] to monitor water surfaces continuously for oil films at critical points on water streams or surfaces. Calculations [327] suggested that reflection of unpolarized light from an oil surface (refractive index n = 1.40) should occur with about 50% greater intensity than from a water surface (n = 1.33) for angles of incidence in the 5–30° range. Experiments with simple
200
Hydrocarbons
laboratory apparatus, i.e. an incandesent lamp, collimating lens, and a photodetector, gave a 100% increase in reflection when a drop of waste oil was added to a water surface. A prototype oil film monitor detected and recorded oil films on a refinery effluent stream flowing at 1ft s-1 (0.3m s-1). Salinity, turbidity and changes in water flow of two to three times had little effect on its behaviour. Operation on a tidal estuary is said to be possible. A baffle was required to eliminate surface turbulence, which is a critical factor, and to avoid fouling. The fouling of glass ‘windows’ on the device was a problem and in the prototype the response to oil was reduced by 50% in 6–8 weeks, although on inland waters weekly cleaning would probably be considered acceptable. Remote detection of oil slicks Mattson [328] has carried out investigations on the application of infrared spectroradiometry to the problem of remote detection and identification of oil slicks, without physical contact with the oil or water. The spectroradiometer incorporates a circular variable interference filter as a monochromator, and a small computer for instrumental control and data reduction. In the preliminary experiments, films of styrene and oleic acid were studied, and spectra between 8.55 and 12.5µm obtained. Radiation was reflected from the oil film, gathered by a Cassegrainian collector, and focused on the circular variable filter wheel of a spectroradiometer, which in these particular experiments allowed scanning between 8.5 and 12.5µm. Thin films of about 10µm thickness tended to produce weakly characteristic styrene reflection spectra, but films of about 300µm produced very definite styrene reflection spectra. The desire to identify the oil, in addition to mere detection, necessitates a complex and even expensive device, unsuitable for general use on inland waters. However, the principle of detection only, based on scanning the reflection over a constant limited range, say 3.3–3.6µm, would be a more practical proposition. Fust et al. [329] have described an instrumental approach to the analysis of oil in water. Ahmadijian and Brown [330] investigated the feasibility of remote detection of water pollutants and oil slicks by laser-excited Raman spectrometry. They showed that, by use of a system of lenses and mirrors, laser-excited Raman spectra can be recorded for samples containing oil at a distance of 21ft (6.4m) from the instrument. McMullen et al. [331] discussed the principal and operation of a system to detect and measure the pollution of water surfaces by oil. The apparatus, which is described in detail with the aid of photographs and drawings, utilizes buoyant sampling heads which float on the water surface to record surface tension changes caused by the presence of either
Mineral oil and petroleum spills
201
soluble or insoluble monolayers. A battery-operated central control unit and pen recorder are attached at the end of a 60ft (18m) floating cable, and a trigger circuit can be used to activate an alarm when surface tension changes are detected. Field trials, using oil as the polluting agent, on various canals are reported, and it is concluded that the apparatus could be used for the detection of detergent and crude sewage pollutants. 2.4.1.8 Preconcentration Adsorption on a molecular sieve 5A is a technique that has been examined for the preconcentration of oil in water samples. Uchiyama [332] gives details of a procedure for the separation and determination of mineral oil, animal oil and vegetable oil in water. After extraction with carbon tetrachloride, the extract is treated with molecular sieve 5A, on which animal and vegetable oils are adsorbed. The oil is then determined by infrared analysis. Carbon disulphide is an excellent solvent in the washing of active carbon traps. An additional advantage of carbon disulphide is the relatively low sensitivity of the flame ionizaton detector for this solvent [333]. Molecular sieves, zeolites and XAD-2 have been used to preconcentrate mineral, animal and vegetable oils from natural waters [97]. 2.4.2 Saline waters 2.4.2.1 Gas chromatography In the marine environment gas chromatography has been employed to identify petroleum products [238, 246, 252, 264]. Here the pollutants are crude oil, marine residual fuel oil and crude oil sludge consisting of a concentrated suspension of high-melting-point paraffin wax in crude oil. Although weathering of marine oil pollutants can be such that the oil is rendered unrecognizable, the time required to achieve this was found to be so long as to be insignificant in regard to pollutant identification. Occasionally the mound of unresolved components on the chromatogram supporting the superimposed n-alkane peaks confuse the true n-alkane profile. This has been overcome by separating off the nalkanes using molecular sieves, prior to gas chromatography [238]. However, separation of n-alkanes in this way, or by urea complex formation [262], is reported as being more applicable to distillates rather than residual materials, and also the separation is not entirely specific [264]. In the case of marine pollutants it has been found advantageous
202
Hydrocarbons
to chromatograph a distilled residue (b.p. > 343°C) [238], or fraction (b.p. 254–370°C) [251], which avoids problems caused by evaporation of lower ends by weathering. Chromatograms of marine pollutants have been published; crude oils, 200s fuel oil, 2000s fuel oil, pollution samples [238]; sludge wax, crude oil sludge, Bahrain fuel oil, 3000s diesel fuel, pollution samples; marine diesel fuel, sludges, crude oil, residual fuel oil [251]. Ramsdale and Wilkinson [258] have identified petroleum sources of beach pollution by gas chromatography. Samples containing up to 90% of sand or up to 80% of emulsified water were identified, without pretreatment by gas chromatography of one of a pair of matched stainless steel columns (750mm × 3.2mm i.d.) fitted with precolumns (100mm) to retain material of high molecular weight, the second column being used as a blank. The column packing is 5% of silicone E301 on Celite (52–60 mesh), the temperature is programmed at 50°C per minute from 50 to 300°C, nitrogen was used as carrier gas, and twin flame ionization detectors were used. Brunnock et al. [238] have also analysed beach pollutants. They showed that weathered crude oil, crude oil sludge and fuel oil can be differentiated by the n-paraffin profile as shown by gas chromatography, wax content, wax melting point and asphaltene content. The effects of weathering at sea in crude oils were studied; parameters unaffected by evaporation and exposure are the contents of vanadium, nickel and nparaffins. The scheme developed for the identification of certain weathered crude oils involves the determination of these constituents, together with the sulphur content of the sample. The Institute of Petroleum [252] in 1970 recommended analytical methods for the identification of the source of pollution by oil of seas, rivers and beaches. Methods are described for recovering the oil from the sample and for the determination of boiling range, the hydrocarbon components are separated in the order of their boiling point by means of gas chromatography. For the determination of metals, after incineration of the oil with sulphur and dissolution of the ash, vanadium and nickel are determined by reaction with tungstophosphate and dimethylglyoxime, respectively, and use of X-ray fluorescence or spectrometry. For the determination of wax content, asphalt-free material is dissolved in hot dichloromethane, the solution is cooled to about -32°C and the precipitated wax is separated and weighed. The fingerprinting of oil spills by gas chromatography has been reported using flame ionization and flame photometric detection [335, 336]. Interpretation of the flame ionization chromatograms is normally based on the distribution of the n-paraffin peaks [334]; however, the use of the pristane-to-phytane ratio has been reported by Blumer and Sass [337] and is thought to be more independent of weathering.
Mineral oil and petroleum spills
203
Boylan and Tripp [338] determined hydrocarbons in sea water extracts of crude oil and crude oil fractions. Samples of polluted seawater and the aqueous phases of simulated samples (prepared by agitation of oil–kerosine mixtures and unpolluted seawater to various degrees) were extracted with pentane. Each extract was subjected to gas chromatography on a column (8ft × 0.06in (2.4m × 1.5mm)) packed with 0.2% of Apiezon L on glass beads (80–100 mesh) and temperature programmed from 60 to 220°C per minute. The components were identified by means of ultraviolet and mass spectra. Polar aromatic compounds in the samples were extracted with methanol– dichloromethane (1:3). McKay [339] has investigated the use of automatic solids injection into the gas chromatographic column in his investigation of hydrocarbon pollution of beach sands. Freegarde [292] has discussed the identification, determination and ultimate fate of oil spilt at sea. Erhardt and Blumer [243] have developed a method for the identification of the source of marine oil spills, by gas chromatographic analysis and results for eight different crude oils are given. Distinguishing compositional features are still recognizable after weathering for more than 8 months. The method was used for the tentative source identification of samples of beach tar. Adlard et al. [336] improved the method of Ramsdale and Wilkinson [258] by using an S-selective flame photometric detector in parallel with the flame ionization detector. Obtaining two independent chromatograms in this way greatly assists identification of a sample. Evaporative weathering of the oil samples has less effect on the information attainable by flame photometric detection than on that attainable by flame ionization detection. A stainless steel column (1m × 3mm i.d.) packed with 3% of OV-1 on AW-DMCS Chromosorb G (85– 100 mesh) was used, temperature programmed from 60 to 295°C per minute with helium (35mL min-1) as carrier gas, but the utility of the two-detector system is enhanced if it is used in conjunction with a stainless steel capillary column (20m × 0.25mm) coated with OV-101 and temperature programmed from 60 to 300°C per minute because of the greater detail shown by the chromatograms. Zafiron et al. [340] have shown that commercial oil spill emulsifiers can interfere with the gas chromatographic detection of the source of oil spill. Investigations on pelagic tar in the North West Atlantic have been carried out by Bulten et al. [341] using gas chromatography. Their report collects together the results of various preliminary investigations. It is in the Sargasso Sea, where the highest concentrations (2–40mg m-2) occur, and on beaches of isolated islands,
204
Hydrocarbons
such as Bermuda. These workers discuss the occurrence, structure, possible sources, and possible fate of tar lumps found on the surface of the ocean. Zafiron and Oliver [342] have developed a method for characterizing environmental hydrocarbons using gas chromatography. Solutions of samples containing oil were separated on an open-tubular column (50ft × 0.02in) coated with OV-101 and temperature programmed from 75 to 275°C at 6°C per minute; helium (50mL min-1) was used as carrier gas and detection was by flame ionization. To prevent contamination of the columns from sample residues, the sample was injected into a glasslined injector assembly, operated at 175°C, from which gases passed into a splitter before entering the column. Analysis of an oil on three columns gave signal-intensity ratios similar enough for direct comparison with a standard. The method was adequate for correlating artificially weathered oils with sources and for differentiating most of 30 oils found in a sea port. Hertz et al. [343] have discussed the methodology for the quantitative and qualitative assessment of oil spills. They describe an integrated chromatographic technique for studies of oil spills. Dynamic headspace sampling and gas chromatography and coupled-column liquid chromatography are used to quantify petroleum samples, and the individual components in these samples are identified by gas chromatography and mass spectrometry. Rasmussen [245] has described gas chromatography methods for the identification of hydrocarbon oil spills. The spill samples are analysed on a 100ft (30.5) Dexsil-300 support coated open tube (SCOT) column to obtain maximum resolution yet retain a high upper temperature limit. The chromatograms are mathematically treated to give ‘gas chromatographic patterns’ that are a characteristic of the oil, yet are essentially unaffected by moderate weathering. Both liquid and solid oil spill samples were analysed with very little sample preparation in less than 2h. Rasmussen [245] compiled the ‘gas chromatographic patterns’ of 20 crude oils. He uses metal and sulphur determinations and infrared spectroscopy to complement the capillary gas chromatographic technique. Rasmussen [245] quotes an example of a bunker oil spill. The spill sample and samples of three potential sources were analysed. Their gas chromatographic patterns were calculated and a comparison confirmed that the oil spill was bunker fuel from a nearby power plant (source A). The chromatograms are shown in Figs 2.15 to 2.18.
Mineral oil and petroleum spills
205
Fig. 2.15 Oil spill sample: C –C segment of chromatogram. (Reprinted with permission from 16 31 Rasmussen [245]. Copyright (1976) American Chemical Society.)
Fig. 2.16 Oil sample from source A: C –C segment of chromatogram. (Reprinted with permission 16 31 from Rasmussen [245]. Copyright (1976) American Chemical Society.)
Gas chromatography–mass spectrometry In some cases it is necessary to identify unambiguously selected components separated during the gas chromatographic examination of oil spill material. Such methods are needed from the standpoint of the enforcement of pollution control laws. The coupling of a mass spectrometer to the separated components emerging from a gas chromatographic separation column enables such positive identifications to be made.
206
Hydrocarbons
Fig. 2.17 Oil sample from source C: C –C segment of chromatogram. (Reprinted with permission 16 31 from Rasmussen [245]. Copyright (1976) American Chemical Society.)
Fig. 2.18 Oil sample from source B: C –C segment of chromatogram. (Reprinted with permission 16 31 from Rasmussen [245]. Copyright (1976) American Chemical Society).
Smith [344] classified large sets of hydrocarbon oil spectra data by computer into ‘correlation sets’ for individual classes of compounds. The correlation sets were then used for determining the class to which an unknown compound belongs from its mass spectral parameters. A correlation set is constructed by use of ion series summation, in which a low-resolution mass spectrum is expressed as a set of numbers representing the contribution to the total ionization of each of 14 ion series. The technique is particularly valuable in the examination of results from coupled gas chromatography–mass spectrometry of complex organic mixtures.
Mineral oil and petroleum spills
207
Walker et al. [345] examined several methods and solvents for use in the extraction of petroleum hydrocarbons from estuarine water and sediments, during an in situ study of petroleum degradation in sea water. The use of hexane, benzene and c h l o ro f o r m a s s o l v e n t s i s d i s c u s s e d a n d c o m p a re d , a n d quantitative and qualitative differences were determined by analysis using low-resolution computerized mass spectrometry. Using these data, and data obtained following the total recovery of petroleum hydrocarbons, it is concluded that benzene or benzene–methanol azeotrope are the most effective solvents. Brown and Huffman [346] reported an investigation of the concentration and composition of non-volatile hydrocarbons in Atlantic Ocean and nearby waters. Sea water samples were taken at depths of 1 and 10m and the non-volatile hydrocarbons were identified by mass spectrometric techniques. The results show that the non-volatile hydrocarbons in Atlantic and nearby waters contained aromatics at lower concentrations than would be expected if the source of the hydrocarbons were crude oil or petroleum refinery products. Hydrocarbons appeared to persist in the water to varying degrees, with the most persistent being the cycloparaffins, then isoparaffins, and finally the aromatics. Albaigés and Albrecht [347] propose that a series of petroleum hydrocarbons of geochemical significance (biological markers) s u c h a s C – C a c y c l i c i s o p re n o i d s a n d C s t e r a n e s a n d 20 40 27 triterpenes are used as passive tags for the characterizaton of oils in the marine environment. They use mass fragmentography of samples to make evident these series of components without resorting to complex enrichment treatments. They point out that computerized gas chromatography–mass spectrometry permits multiple fingerprinting from the same gas chromatographic run. Hence rapid and effective comparisons between samples and long-term storage of the results for future examination can be carried out. Usually, Albaigés and Albrecht [347] first deasphaltenized with n-pentane (40 volumes) prior to gas chromatography. When, however, the recovery of the branched plus cyclic alkanes was needed for subsequent analysis the saturated hydrocarbon fraction was isolated by conventional silica gel adsorption chromatography (eluting solvent n-pentane) and refluxed in isooctane with 5Å (0.5nm) molecular sieves. Gas chromatographic profiles of petroleum residues, one example of which is shown in Fig. 2.19, exhibit several characteristic features that have been applied for identification or correlation purposes.
208
Hydrocarbons
Fig. 2.19 FID and FPD high-resolution chromatograms of a crude oil residue (b.p. > 220°C). Numbers above the peaks (with or without asterisk) indicate the number of carbon atoms of the isoprenoid and the n-paraffin hydrocarbons, respectively. (Reprinted with permission from Albaigés and Albrecht [347]. Copyright (1979) Gordon and Breach.)
Generally, the most apparent is the n-paraffin distribution (upper trace in Fig. 2.19) that has proved to be useful in differentiating the main types of pollutant samples (crude oils, fuel oils and tank washings) [258] or even types of crude oils [348], although in this case the method involves the quantification of the previously isolated n-paraffins, therefore lengthening the analysis time. Another relevant feature of the gas chromatographic profile is the acyclic isoprenoid hydrocarbon pattern that is made evident with capillary columns (peaks with an asterisk in the upper trace of Fig. 2.19) or by inclusion of the saturated fraction in 5Å (0.5nm) molecular sieves or in urea (middle trace in Fig. 2.19). The predominant peaks usually correspond to the C (pristane) and C (phytane) isomers, the 19 20 ratio of which serves as an identification parameter [243], although the series extends to lower and higher homologues.
Mineral oil and petroleum spills
209
Finally, the sulphur compounds that are present in minor quantities in petroleum products also exhibit a typical gas chromatographic fingerprint easily obtained by flame photometric detection (FPD, lower trace in Fig. 2.19). This fingerprint has been introduced to complement the flame ionization detection (FID) chromatogram with the aim of resolving the ambiguities or increasing the reliability in the identification of the pollutants [336]. All the above fingerprints exhibit a different usefulness for characterizing oils. Their variation between crudes and their resistance to the sea-weathering process are not enough, in many cases, for providing the unequivocal identification of the pollutant. The n-paraffins can, apparently, be removed by biodegradation as well as the lower acyclic isoprenoids at respectively slower rates [349]. On the other hand, fractions boiling at up to 300°C can be lost by evaporation affecting both n-paraffin and the isoprenoid distributions. The flame photometric detector chromatogram is less sensitive to modification by bacterial metabolism but can also be affected by evaporation, in spite of its higher retention range (Fig. 2.19), as will be shown later. However, the last part of the flame ionization detector chromatogram (Fig. 2.19) appears to be highly promising in overcoming these limitations. In fact, this part corresponds to a hydrocarbon fraction that boils at over 400°C, so it cannot easily be evaporated under environmental marine conditions. Moreover, it contains a wealth of compounds of geochemical significance, namely isoparaffins and polycyclic alkanes of isoprenoid, sterane and triterpane structure [350, 351] as a result of a complete reduction of precursor isoprenyl alcohols, sterols and triterpanes, respectively. Therefore, their occurrence and final distribution in crude oils will be related to their particular genetic history, that is to the original sedimentary o rganic matter and to the processes undergone during its geochemical cycle. In consequence, it was assumed by Albaigés and Albrecht [347] that these factors can provide unique hydrocarbon compositions for each crude oil, by which the unambiguous identification of the samples can be brought about. Besides their geochemical stability, these compounds do also remain unaltered after biodegradation [352], being, in this respect, valuable passive tags for characterizing marine pollutants. The problem is that such components are present frequently at trace levels, as part of very complex mixtures and can only be recognized after long and tedious enrichment treatments that are not practical from the standpoint of the routine or monitoring analysis. Mass fragmentography provides a satisfactory tool for obtaining specific fingerprints for classes and homologous series compounds,
210
Hydrocarbons
resolved by gas chromatography. In addition, computerized gas chromatography–mass spectrometry allows multiple fingerprinting from the same chromatogram, which is especially important for a quick survey of any compound class in a scanty sample and permits storing the information for further processing or correlation studies. However, to carry out the analysis successfully a precise knowledge of the nature and the gas chromatographic and mass spectrometric behaviour of such compounds is needed. Albaigés and others have done considerable work in recent years on this topic [353–356]. 2.4.2.2 Paper chromatography One of the first reported applications of paper chromatography to the identification of petroleum pollutants was that of Schuldiner [357] who was able to distinguish between crude oil, marine fuel oil and crude oil sludge. A variation of paper chromatography occasionally referred to as evaporation chromatography was employed by Herd [358] to compare fuel oils suspected of being identical. Paper strips were immersed in an ether solution of the oil and the ether allowed to evaporate, usually overnight. Characteristic bands were revealed under ultraviolet light. 2.4.2.3 Gel permeation chromatography Done and Reid [359] applied gel permeation chromatography to the identification of crude oils and products isolated from estuary and sea water. The technique, which appears more suited to the analysis of crude oils, is based on the separation of oil components in order of their molecular size, for practical purposes their molecular weight. Oils were dissolved in tetrahydofuran or toluene and each solvent resulted in a different output profile, and therefore more information for identification. The solutions were pumped through a 24in × 3/8in (0.6m × 9.5mm) column packed with 6nm (60Å) Styragel and the eluant monitored by a differential refractometer. Elution time was 1h, although this was improved by staggered injection, using 6mg samples. ‘Fingerprints’ of crude oils investigated fall into several discrete groups. Crude oil residues (b.p. 525°C) were excluded by the Styragel employed, while crude oil and weathered samples were found to give similar traces. Some changes were introduced by evaporation processes, and it could be difficult to differentiate between similar but weathered crude oil. Heavy fuel oils were more difficult to identify, owing to the variety of fuel oils in use.
Mineral oil and petroleum spills
211
2.4.2.4 Infrared spectroscopy Kawahara [75, 240, 246, 259] has described an infrared spectroscopic method applicable to essentially non-volatile petroleum products. The author states that useful information cannot be obtained from infrared spectroscopy applied to volatile products such as naphthas and gasolines, unless on-the-spot collection of the sample is made immediately following a spill [75]. Heavy residual fuel oils and asphalts are not amenable to gas chromatography and give similar infrared spectra. However, a differentiation can be made by comparing certain absorption intensities [75]. Samples were extracted with chloroform, filtered, dried, and the solvent evaporated off at 100°C for a few minutes using an infrared lamp. A rock salt smear was prepared from the residue in a little chloroform, and the final traces of solvent removed using the infrared lamp. The method, which in effect compares the paraffinic and aromatic nature of the sample, involves calculation of the following absorption intensity ratios: • • • • • •
(13.88µm polymethylene chain)/(7.27µm methyl groups); (3.28um aromatic C–H)/(3.42µm aliphatic C–H); (12.34µm aromatic rings)/(7.27µm methyl groups); (6.25µm aromatic C–C)/(7.27µm methyl groups); (12.34µm aromatic rings)/(13.88µm polymethylene chain); (6.25µm aromatic C–C)/(13.88µm polymethylene chain).
Peaks observed at 5.90µm and 8.70µm were thought to reflect oxidative effects on the asphaltic material, while asphaltic sulphoxide and sulphone were tentatively inferred by bands at 9.76, 8.66 and 7.72µm. The 12.34/ 13.88, 12.34/7.27 and 6.25/13.88µm ratios tended to show the greatest difference between different samples. When the ratio 12.34/7.27µm versus 12.34/13.88 were plotted graphically, the intermediate fuel oils behaved similarly [240]. Weathering caused fuel oils to fall below the curve although with asphalts the effect was significantly less. Since no prior purification was employed the method relies on an uncontaminated, unweathered sample of oil being available. Mattson and Mark [360] reported some criticism of Kawahara’s technique. They claim that evaporation of the solvent chloroform by infrared heating removes volatiles and causes large changes in the ratios. An oil sample was shown to suffer such alteration by the infrared during repeated analysis. The absorption of all bands decreased non-uniformly between 20 and 100% over a period of 30min. They propose the application of internal reflection spectrometry as a rapid, direct qualitative technique requiring no sample pretreatment. Unlike infrared spectrometry which involves transmission of infrared radiation through a homogeneous transparent solution,
212
Hydrocarbons
infrared reflection spectrometry is based on attenuated reflection of the infrared radiation between the sample layer and a suitable, highly polished crystal surface. The attenuation of the radiation passing through a cell is monitored and recorded in the same manner as transmission infrared spectrometry, although the resulting spectrum shows absorptions at slightly shifted wavelengths [361]. In contrast to infrared spectrometry there is no decrease in relative sensitivity in the lower energy region of the spectrum, and since no solvent is required, no part of the spectrum contains solvent absorptions. Oil samples contaminated with sand, sediment and other solid substances have been analysed directly, after being placed between 0.5mm 23-reflection crystals. Crude oils, which were relatively uncontaminated and needed less sensitivity, were smeared on a 2mm 5-reflection crystal. The technique has been used to differentiate between crude oils from natural marine seepage, and accidental leaks from a drilling platform. The technique overcomes some of the faults of infrared spectroscopy, but is still affected by weathering and contamination of samples by other organic matter. The authors suggested the absorption bands shown in Table 2.15 as being important in petroleum product identification. Kawahara (Kawahara and Ballinger [240], Kawahara [271]) has used his method to characterize a number of known and unknown petroleum samples. All of these studies used the normal transmission method to obtain infrared spectra; however, the feasibility of using internal reflection to obtain infrared spectra has been demonstrated by several groups (Mattson and Mark [360], Mark et al. [362] and Baier [363]). The advantage of the latter method is that chemical extraction of petroleum from such as sand and water is unnecessary. Table 2.19 Absorption bands important in petroleum product identification [361]
Mineral oil and petroleum spills
213
Mattson [364] has also reported on investigations of samples from 40 oil sources and has shown that there exist sufficient differences between the fingerprints by infrared spectroscopy of oils from various sources such as oil spills for the technique to be used to identify them. Lynch and Brown [241] have obtained infrared spectra for over 50 samples of crude oils, fuel oils and other petroleum products. These were characteristic of each sample. They showed that computer analysis of absorptivities can be used to match unknown samples with those already known. Lynch and Brown are of the opinion that with appropriate data analysis infrared spectroscopy can be used to identify the type and source of a reasonably large number of petroleum samples. The method of Lynch and Brown [241] is an extension of the ones suggested by Kawahara [75] and Mattson [364]; however, they used additional low-frequency bands and higher instrument resolution. Furthermore, they developed a new method for numerical analysis of the spectra data, which utilizes a digital computer to match an unknown petroleum sample with the correct known. These workers used a high resolution instrument. The analysis of Lynch and Brown [241] both by visual inspection and on a computer is confined to bands from 650 to 1200cm-1. Other investigators have used bands in this region, e.g. Kawahara [75] used bands at 720 and 810cm-1, whereas Mattson and Mark [360, 361] used bands at 725, 747, 814 and 1034cm -1 . Spectra for these previous investigations were measured with lower resolution. Using a highresolution instrument Lynch and Brown [241] observed many sharp bands between 650 and 1200cm-1; this is especially true in the 700– 900cm-1 region where they observed as many as 19 bands in spectra of crude oils and the lighter distillates. The intensities and the number of bands between 650 and 1200cm -1 are extremely characteristic of petroleum samples, and they provide a unique fingerprint of each individual sample. The fingerprint can be used to match an unknown sample to the correct known rapidly and unambiguously. Lynch and Brown [241] investigated the applicability of computer analysis of infrared spectral data on unknown oil samples to the identification of the origin of the oil. Lynch and Brown [241] quote several examples of the application of their technique to the identification of oil spill materials. They claim that it provides an unambiguous identification for petroleum products, eliminating the need for adding tracer materials to petroleum products or for other methods of analysis. Ideally, for the method to be used on a worldwide scale, infrared spectra of all possible petroleum samples would have to be measured and the absorptivities of the selected bands stored in a computer file. This could all be accomplished within a relatively short period of time. In addition, to identify samples taken
214
Hydrocarbons
from natural waters, the effects of weathering on the infrared spectra of samples would have to be known. Ahmed et al. [365] investigated the precision and accuracy of infrared spectroscopic methods for determining oil in estuary water using two techniques of oil preconcentration, namely carbon tetrachloride extraction and polyurethane foam adsorption. The average recovery is 101% and the average error is 13%. Therefore, the method itself seems quite reliable since the quality of oil in the sample bottle can be determined with a relative standard deviation of less than 15% for concentrations exceeding about 0.055mg L-1. 2.4.2.5 Raman spectroscopy Ahmadjian and Brown [366] have used laser Raman spectroscopy to identify petroleums. 2.4.2.6 Ultraviolet spectroscopy The absorption of ultraviolet radiation is much less useful than fluorescence techniques. Spectra of materials as complex as petroleum products are insufficiently detailed for identification purposes. Levy [125] has identified petroleum products in a marine environment by ultraviolet spectroscopy. The sample (down to 0.06mg) in hexane is filtered through Whatman No. 42 paper, the filtrate is diluted to 1000mL and the extinction is measured at 256nm. The dilution is adjusted, if necessary, to bring the excitation within the range 0.4–0.8, and the spectrum relative to hexane is then recorded over the range 350– 210nm. The absorption at about 256nm and about 228nm is measured, the ratios of these values is independent of concentration of the test solution and can be used for distinguishing between samples of residual fuel oils and lubricating oils even after weathering. 2.4.2.7 Fluorescence spectrometry Fluorescence spectrometry has been adapted as an alternative analytical method for estimating oil in various environmental samples. The method was originally used to detect oil in surveys of oil in sea water (Zitko and Carson [367]; Michalik and Gordon [368]; Levy [369, 370]; Levy and Walton [371]). Hargrave and Phillips [372] have used fluorescence spectrometry to evaluate concentrations of aromatic constituents in aquatic sediments, using excitation at 310µm and measuring the 374nm emission. The oil concerned, a Venezuelan crude, contained about 35% by weight of aromatic constituents. Aromatic substances were extracted with n-
Mineral oil and petroleum spills
215
hexane and fluorescence spectroscopy was used to produce a series of contour diagrams of fluorescence intensity at various excitation and emission wavelengths, in order to compare fluorescence spectral patterns of sample extracts and standard oils. Petroleum residues were determined and it was found that total oil concentrations ranged from 10 to 3000µg g-1 wet sediment, with the highest concentrations occurring in sedimenting particles. Standardized spiking experiments were performed by Hargrave and Phillips [372] with various amounts of Venezuela crude oil (approximately 35% aromatic compounds by weight) added to freshly collected beach sand and mud sediments in various concentrations. Recovery efficiencies of total oil added with the n-hexane extractant were always in excess of 95%. Several experiments were conducted to determine the optimum hexane/ water ratio to be used for extraction. When no distilled water was present, wet sediment coagulated in hexane and fluorescence intensity in extracts was reduced in comparison with that achieved with water present. The addition of distilled water produced a slurry of sediment and hexane and the fluorescence intensity in extracts was increased. Water/hexane volume ratios between 0.1 and 0.2 resulted in maximum extraction of fluorescing compounds and minimized variance between replicate samples. Hargrave and Phillips [372] point out that application of the fluorescence method for estimating oil in water, sediments and organisms requires that fluorescence be related to a standard. Wavelengths for measurement were chosen by scanning excitation and emission spectra and identifying peaks. An oil which contains similar excitation and emission maxima is then used as a standard to produce a calibration. Thus, total oil concentrations are not measured, only inferred, and the use of such calibration assumes that all fluorescence in samples is attributable to the presence of fluorescing compounds equivalent to those in the petroleum standard. Parker and Barnes [297] and Lloyd [294–296] have demonstrated that fluorescence spectral analysis, using combinations of excitation and emission wavelengths, can be employed to characterize mixtures of aromatic compounds. Freegarde et al. [373] suggest that complete variations in fluorescence spectral patterns can be represented if emission spectra are scanned over a range of different excitation wavelengths and contours of equal fluorescence plotted. Such contour diagrams topographically indicate the combination of excitation and emission wavelengths which results in maximum fluorescence and permits comparison of fluorescence patterns produced by the aromatic substances present in different samples. Freegarde et al. [373] extended the technique to crude oil in sea water, which was extracted by cyclohexane and the fluorescence emission
216
Hydrocarbons
spectra measured. The sensitivity was limited by the blank. The minimum detectable oil content was about 0.001mg L-1. 2.4.2.8 Characterization of oils by metals analysis Lieberman [374] discussed the application of various techniques of metals analysis to oil pollution source identification. He subjected crude oils, residual fuel oils, and distillates fuel oil to simulated weathering for 10 to 21 days, at 55°F (13°C) and 80°F (27°C), at high and low rates of salt water washing. ‘Weathered’ and ‘unweathered’ samples were analysed by lowvoltage mass spectroscopy (for polycyclic aromatics), high-voltage mass spectroscopy (naphthenes), gas chromatography (n-alkanes), emission spectroscopy (Ni and V), X-ray spectroscopy (total S) and the Kjeldahl technique (total N). Several compound indices were sufficiently unaffected by simulated weathering to allow discrimination between like and unlike pairs of oils. Discriminant-function analysis was used to select the best compound indices for the oils under examination. By using these indices, weathered and unweathered samples were correctly paired with high statistical confidence. Various other workers have investigated methods for the identification of oil spills [334–337]. Wilson et al. [334] reviewed approaches based on trace metal (Ni/V) ratio, nitrogen and sulphur content, infrared spectroscopy and gas chromatography. 2.4.2.9 Miscellaneous Remote detection of oil spills Millard and Arvesen [375] discuss the airborne optical detection of oil on water. They undertook absolute radiometry, differential radiometry and polarimetry measurements utilizing reflected sunlight over the range 380–950nm to evaluate methods for detecting oil spills on sea water. Maximum contrast between oil and water was observed at wavelengths less than 400nm and greater than 600nm, minimum contrast being in the range 450–500nm. Oil always appeared brighter than the water, but it was not possible to distinguish one oil from another. These workers commented that differential polarization appeared to be a promising technique. Pierre [376] has reported an exploratory study of the characterization of surface oil slicks by infrared reflective spectroscopy. A double-beam spectrophotometer was modified for studying the reflectance spectra (at angles of incidence of 45°, 60° and 70°) of oil layers (20–30µm thick) on the surface of water using pure water as reference.
Mineral oil and petroleum spills
217
Freegarde [292] has reviewed methods of identifying and determining oils in sea water. They describe a method for determining down to 0.001mg L-1 of crude oil in sea water. 2.4.3 Waste waters 2.4.3.1 Gas chromatography Cole [72] has investigated the use of gas chromatography in the identification of slop oils resulting from oil refinery leaks. Oils leaking into the waste water system were examined by gas chromatography on a column (6ft × 0.25in o.d. (1.8m × 6mm)) of 20% of SE-52 on silanized Chromosorb W (60–80 mesh) temperature programmed from 50 to 300°C at 15°C per minute, with nitrogen (60mL min-1) as carrier gas and flame ionization detection, and by infrared spectrophotometry. Reference is then made to a library of the results of similar tests on samples of all the refinery process streams. Only 1mL of sample is required, and identification normally takes less than 1h. Martin and Geyer [377] also studied infrared spectroscopy as a means of estimating mineral oils in water. The method was unsatisfactory in the case of waste waters containing emulsifying agents, which must be removed, together with other polar compounds, by extraction with carbon tetrachloride and treatment of this extract with an adsorption agent. They compared results obtained by shaking with Florasil or by passing through a chromatographic column of aluminium oxide or Florasil. For some waste waters, containing phenol, only the aluminium oxide column can be used. It is recommended that the Florasil used for adsorption of polar compounds should be replaced by aluminium oxide. 2.4.4 Trade effluents 2.4.4.1 Fluorescence spectrometry Most published procedures deal with the determination of petroleum products in industrial effluents from flotation processes, boilers, refineries etc. [378–389]. Usually the oil is extracted from the aqueous sample with a small volume of organic solvent, for example, benzene, petroleum ether [378, 384, 385], chloromethane, diethyl ether, or toluene [387] or nonfluorescing gasoline [384]. In all these methods, a sample of the polluting oil was available and hence calibration was possible. Often the fluorescence level of the blank was that of the incoming process or boiler feed water. The intensity of fluorescence was determined in commercial
218
Hydrocarbons
fluorimeters, and concentration limits of 1.5–0.2mg L-1 [382], 100–300mg L-1 with ±0.7% error [378], and a lower detection limit of 0.5µg L-1 [329] have been reported. 2.4.5 Sewage effluents 2.4.5.1 Miscellaneous Millson [390] investigated components of sewage sludge and found elementary sulphur in the hydrocarbon fractions eluted from liquid adsorption columns. By using a solid adsorbent such as alumina, silica gel, or Florasil, and heptane as eluant, the sulphur could be separated from weakly adsorbed hydrocarbons, e.g. squalane or biphenyl, but not from more strongly adsorbed hydrocarbons such as phenyldodecane. 2.5 Mineral greases The traditionally and widely accepted method for determining grease in waste water is based on organic solvent extraction, solvent evaporation and weighing [88]. The method is time-comsuming, both in the analyst’s time and in duration. A rather large sample of waste water is required for the traditional method and it is difficult to manipulate the samples to assure a homogeneous aliquot. Simplification of the methodology is fraught with difficulties because of the extreme complexity of waste water in composition and variability. Taras and Blum [391] have pointed out that the scope of the standard method of the American Public Health Association for the determination of grease can be extended to include emulsifying oils by saturating the acidified sample with sodium chloride before filtration. Trichlorotrifluoroethane and hexane are equally effective as extracting solvents, but the former is preferred on account of its noninflammability and greater density. The most difficult oil sample studied was about 85% recoverable by the modified method. In view of the unsatisfactory situation regarding gravimetric methods for the determination of oil in water it is recommended that alternative, more specific, techniques are considered as discussed below. Maxcy [392] has described improved methods for the determination of grease in waste water. Cook et al. [393] have described improved methods for the determination of oil and grease in water. This method utilizes the semiautomatic oil monitoring instrument, the Horiba OCMA-200. These workers compared results obtained by this procedure with those obtained by a US Environmental Protection Agency reference method
References
219
[394]. The Horiba instrument is a very convenient device to use since sample handling and contamination possibilities are almost nil. An acidified, 10mL oil-in-water sample is injected via a syringe into an extraction chamber with an equal amount of solvent, either Freon or carbon tetrachloride. A vibrating plunger is activated for a given time causing intimate mixing of the sample and solvent. The plunger is deactivated and the solvent and oil mixture is allowed to separate. A valve is then opened, permitting the solvent mixture to flow through a filter into the infrared chamber where the amount of 3.5µm light attenuation by the oil in solvent is measured. Cleverley [398] has carried out a comparison of various lubricating greases by infrared spectroscopy in the 4000–650cm-1 region. This work would be useful in classifying greases originating in extracts of water samples. Various workers [395–397] have discussed the volumetric determination of greases in sewage, sludges and industrial wastes. 2.6 Vegetable oils The Standing Committee of Analysts [399] compared oils and grease determinations in water obtained by solvent extraction–infrared spectroscopy and by solvent extraction gravimetric methods. In the case of the gravimetric method there was minimal boiling limitation on the materials determined. Oils and fats of natural origin are retained on the Florasil column employed in the initial separation and may be determined separately by elution with a suitable solvent. The National Water Council (UK) [400] have described a petroleum ether extraction procedure for the determination of oils and fats in waste water. References 1 2 3 4 5 6 7 8 9 10 11 12
Lively, L. Engng. Bull. Ext. Serv. Purdue University 118 657 (1965) Jeltes, R. and Veldink, R. Journal of Chromatography 27 242 (1967) McAucliffe, C. Journal of Physical Chemistry 70 1267 (1966) Bridie, A. L., Bos, J. and Herzberg, S.J. J. St. Petrol. 59 263 (1973) Desbaumes, E. and Imhoff, C. Water Research 6, 885 (1972) Khazal, W.J., Vejostra, J. and Novák, J. Journal of Chromatography 157 125 (1978) Drodz, J. and Novák, J. Journal of Chromatography 152 55 (1978) Drodz, J. and Novák, J. Journal of Chromatography 136 37 (1977) Drodz, J. and Novák, J. Journal of Chromatography 158 471 (1978) McAucliffe, C. Chemical Technology 1, 46 (1971) Kaiser, L.E. and Oliver, B.G. Analytical Chemistry 48 2207 (1976) McAucliffe, C. Journal of Chemical Geology 4 225 (1969)
220
Hydrocarbons
13 Swinnerton, J.W. and Linnenbom, V.J. Journal of Gas Chromatography 5 570 (1967) 14 Novák, J., Zlutický, J., Kubelka, V. and Mostecký, J. Journal of Chromatography 76 45 (1973) 15 Grob, K. Journal of Chromatography 84 255 (1973) 16 Grob, K. and Zurcher, F. Journal of Chromatography 117 285 (1976) 17 Grob, K. and Grob, G. Journal of Chromatography 90 303 (1974) 18 Grob, K., Grob, G. and Grob, K. Journal of Chromatography 106 303 (1975) 19 Polak, J. and Lu, B. Anal. Chim. Acta 63 231 (1974) 20 Kaiser, R. Journal of Chromatographic Science 9 227 (1971) 21 Schönmann, M. and Kern, H. Varian Instrument News 15 6 (1981) 22 Colenutt, B.A. and Thorburn, S. International Journal of Environmental Studies 15 25 (1980) 23 Drodz, J., Vodakova, Z. and Novák, J. Journal of Chromatography 354 47 (1986) 24 Jeltes, R. Water Pollution by Oil (editor P. Hepple), London, Institute of Petroleum, p. 43 (1971) 25 Simard, R.C., Hasecawa, W., Boudaruk, W. and Headington, C.E. Analytical Chemistry 23 1384 (1951) 26 Ruebelt, C. Helgoländer Wiss. Meeresunters. 16 306 (1967) 27 Fastabend, W., Chemie-Ingr.-Tech. 37 728 (1965) 28 Ruebelt, C. Gas- u. Wass. Fach. 108 893 (1967) 29 Ruebelt, C.Z. Analytical Chemistry 221 299 (1966) 30 Osipov, V.M. Khimiya Tekhnol. Topl. Masel. 16 52 (1971) 31 Galubeva, M.T. Lab. Delo. 11 665 (1966) 32 Lindgreen, C.G. Journal of American Water Works Association 49 55 (1957) 33 Ludzack, F.J. and Whitefield, C.E. Analytical Chemistry 28 157 (1956) 34 Benyon, L.R. Methods for the Analysis of Oil in Water and Soil, The Hague, Striching CONCAWE 40 pp. (1968) 35 Hughes, D.R., Belcher, R.S. and O’Brien, E.J. Bull. Eviron. Contam. Toxicol. 10 170 (1973) 36 Suzuki, R., Yamaguchi, N. and Matsumoto, N. Japan Analyst 23 1296 (1974) 37 Rosen, A.A. and Middleton, F.M. Analytical Chemistry 27 709 (1955) 38 Webber, L.A. and Burks, C.E. Analytical Chemistry 24 1086 (1952) 39 Santo, T., Hagiwara, K. and Ozawa, Y. Japan Analyst 21 1235 (1972) 40 Brown, R.A., Searl, T.D., Elliot, J.D., Phillips, B.G., Brandon, D.E. and Horoghan, H. Proc. of 1973 Conference on Prevention and Control of Oil Spills, Washington DC p. 505 (1973) 41 Rodier, J. Chemical and Physicochemical Analysis of Water, Paris, Dunod, 700 pp. (1971) 42 Boisselet, L. Proc. 5th World Petroleum Congress, New York, pp. 165–175 (1959) 43 Hellmann, H.Z. Anal Chem. 269 245 (1973) 44 Gruenfeld, M. Environmental Science and Technology 1 636 (1973) 45 Mallevialle, J. Water Research 8 1071 (1974) 46 Götz, R. Facum Städte Hygiene, 29 10 (1978) 47 Powell, D.L., Darland, E.J. and Williams, T.R. Analytical Letters 4 479 (1971) 48 Golden, J. Techniques et Sciences Municipals 71 17 (1976) 49 Liu, D.L. Water and Sewage Works 125 40 (1978) 50 Whittle, P.J., McCrum, W.A. and Horne, M.W. Analyst (London) 105 679 (1980) 51 Stichling CONCAWE, The Hague, Report 9/72 (1972)
References
221
52 US EPA Manual of Methods for Chemical Analysis of Water and Wastes, p. 232, Storet No. 00560 (1974) 53 American Petroleum Institute. Manual of Disposal of Refinery Waste, Vol. IV, Method 733-58 (1958) 54 Coles, G.P., Dill, R.M. and Shull, D.L. Am. Chem. Soc., Div. Pet. Chem. Prep. 20 641 (1975) 55 Berthold, I.Z.Z. Anal. Chem. 240 320 (1968) 56 Koppe, P. and Muhle, L.A. Vom Wasser 35 42 (1968) 57 Hunter, D. Environmental Science and Technology 9 241 (1975) 58 Sinel’nikov, V.E. Gidrokhim. Mater. 50 127 (1969). Ref. Zhur. Khim. 19GD (1969) (24) Abstract No. 249.264 59 Legrande, A., Capdeville, H., Rogues, H. and Hoornaert, P. Tribune de Cebedeau 38 19 (1985) 60 McAucliffe, C.D. Chem. Engng. Progress 83 40 (1987) 61 Gibb, J.P. and Barcelona, M.J. Journal of American Water Works Association 78 60 (1986) 62 Pankow, J.F., Isabelle, L.M., Hewertson, J.P. and Cherry, J.A. Groundwater 22 230 (1984) 63 Fam, S., Stenstrau, M.K. and Silverman, G. J. Environmental Engineering 113 1032 (1987) 64 Wade, T.L. and Quinn, J.G. Marine Pollution Bulletin 6 54 (1975) 65 May, W.E., Chester, S.N., Cram, S.P., Gump, B.H., Hertz, H.S., Evagonio, D.P. and Dryzel, S.M. Journal of Chromatographic Science 13 535 (1975) 66 Zsolnay, A. and Kiel, W.J. Journal of Chromatography 90 79 (1974) 67 Witmer, F.E. and Goilan, A. Environmental Science and Technology 7 945 (1973) 68 Matsumoto, G. and Hanya, T. Water Research 15 217 (1981) 69 Rudge, R. and Nicholas, P.V.T. Ass. Analysts 8 31 (1970) 70 Rosen, O. Severn River Authority, private communication 71 Voege, F.A. and Vanesche, H.L. Journal of American Water Works Association 56 1351 (1964) 72 Cole, R.D. Journal of the Institute of Petroleum (London) 54 288 (1968) 73 Machler, C.Z. and Greenberg, A.E. Sanit. J. Engng. Div. Am. Soc. Civ. Engrs. 94 969 (1968) 74 Bogatie, C.F. Tappi 57 103 (1974) 75 Kawahara, F.K. Environmental Science and Technology 3 150 (1969) 76 Kawahara, C.F. and Ballinger, D.C. Ind. Engng Chem. Prod. Res. Dev. 4 553 (1970) 77 Rotteri, S. International Journal of Environmental Analytical Chemistry, 11 263 (1982) 78 Geobgen, H.G. Haustech Essen, Vortragsveroeff. 231 55 (1970) 79 Goretti, G., Morsella, I., and Petronio, B.U. Rio Ital. Sostanze Grasse 51 66 (1974) 80 Strensom, M.K., Fam, S., and Silverman, G.S. Environmental Technology Letters 7 625 (1986) 81 American Public Health Association. Standard Methods for the Examination of Waters and Wastewaters (1985) 82 Lee. E.G.H., and Walden, C.C. Water Research 4 641 (1970) 83 Association of British Chemical Manufacturers and Society for Analytical Chemistry. Recommended Methods for the Analysis of Trade Effluent, Cambridge, Heffer, p. 1244 (1958) 84 Milner, A. Private communication 85 Sherratt, J.G. Analyst (London) 81 518 (1956)
222
Hydrocarbons
86 Rogen, J. Chemical and Physicochemical Analysis of Water, Paris, Dunod, p. 700 (1971) 87 American Society for Testing and Materials Standards with Related Materials, Part 23, Water and Atmospheric Analysis, Philadelphia, The Society, XVIII, 1036 pp. (1969) 88 American Public Health Association. Standard Methods for Examination of Water and Wastewater, 13th Edition, New York, The Association, XXXV, 874 pp. (1971) 89 The Oil Companies International Study Group of Conservation of Clean Air and Water, Europe CONCAWE report no. 6/82 , Analysis of trace substances in aqueous effluents from petroleum refineries (1982) 90 Environmental Protection Agency Report EPA 624, Environmental Protection Agency, U.S.A. 91 Environmental Protection Agency Report EPA 602, Environmental Protection Agency, U.S.A. 92 Environmental Protection Agency Report EPA 601, Environmental Protection Agency, U.S.A. 93 Copin. A., Delen, R., Monteau, A. and Rocher, M. Science of the Total Environment 22 179 (1982) 94 Bohne-Mabusall, R., Otto, K. and Wilderer, P. Z. Wasser Abwasser Forsch. 20 35 (1987) 95 Department of the Environment and National Water Council, HM Stationery Office, London. Methods for the Examination of Water and Associated Materials, 12 pp. (R22 BEN) (1979) 96 Roberts, A.J. and Thomas, T.C. Environmental Toxicol. Chem. 5 3 (1986) 97 Belkin, F. and Hable, M.A. Bulletin of Environmental Contamination and Toxicology 40 244 (1988) 98 Ruebelt, C. Haustech Vortragsveroeff. 65 231 (1970) 99 Sememov, A.D., Stradomanskaya, A.G. and Zurbina, L.F. Gidrokhim. Mater. 53 51 (1971) Ref. Zhur. Khim. 199D. (1971) 100 Bundt, J., Herbel, W. and Steinhart, H. Fresenius Z. Analyt. Chemie 328 480 (1987) 101 Wasik, S.P. Journal of Chromatography 12 845 (1974) 102 Shtivel, N.K., Gipokova, L.I. and Smirnova, Z.S. Soviet Journal of Water Chemistry and Technology 7 66 (1985) 103 Reisus, K. Fortsche. Waterchem. Ihrer, Grenzeb. 10 43 (1968) 104 Mallevialle, J., Lefevre, M., Roussau, C. and Sagbes, C. Rev. Franc Scil. 1 125 (1982) 105 Wasik, S.P. and Tsang, W.W. Analytical Chemistry 42 1649 (1970) 106 Wasik, S.P. and Tsang, W. Analytical Abstracts 21 1854 (1971) 107 Mel’kanovitskaya, G.C. Gidrokhim. Mater. 53 153 (1972). Ref. Zhur. Khim. 199D (1972). Abstract No. 129265 108 Stolyarov, B.V. Chromatographia 14 699 (1981) 109 Urano, K., Meada, H., Ogura, K. and Wada, H. Water Research 16 323 (1982) 110 Senn, R.B. and Johnson, M.S. Ground Water Monitoring Review 7 58 (1987) 111 Dudova, M.Ya. and Diterikhas, O.D. Gidrokhim. Mater. 50 115 (1969) Ref. Zhur. Khim. 19GD (1969) (23) Abstract No. 23G246 112 Geyer, D., Martin, P. and Adrian, P. Gas- u. Wasserfach. Wasser Abwasser 119 72 (1978) 113 Louch, D., Motlagh, S. and Paliszgn, J. Analytical Chemistry 64 1187 (1992) 114 Imanvilov, L.A. Transp. Khranenie Nefteprod. Uglevovdorodn Syr’ya 4 7 (1986) 115 Saltzman, R.S. Analysis Instrum. 6 79 (1968)
References
223
116 Obnorlenskii, P.A. Khimiya Tekhnol. Topl, Masel. 15 54 (1970) 117 Nadzhafova, K.N. In Russian, Trudy vses. Nauchno-issled. Inst. Vodosnabzh., Kanaliz., Gidtotekhn. Sooruzhenii Inzh. Gidrogeol., No. 23, 1007 (1970) 118 Minasyan, K.V. In Russian, Prom. Arm. 11 50 (1969) 119 Ospimov, V.M. and Belova, T.D. In Russian, Khimiya Tekhnol. Topl. Masel. 13 56 (1968) 120 Skotnikova, L.A.. In Russian, Izv. Vyssh. Ucheb. Zaved., Neft. Gaz. 12 111 (1969) 121 Polinskaya, R.E. In Russian, Proekt. Issled. Rab., Neftedob. Prom., Giprovostokneft No 3, 106 (1967) 122 Lurje, Yu.Yu. In Russian, Nauch. Soobshch. Veses. Nauchno-issled. Inst. Vodosnabzh., Kanaliz., Gidrotekhn. Sooruzhenii, Inzh. Gidrogeol., Ochistka Prom. Stochn. Vod. Moscow 34 (1963) 123 Harva, O. and Somersalo, A. Acta Chem. Fenn. 31 384 (1958) 124 Przybylski, Z. Chemia Analit. 8 601 (1963) 125 Levy, E.M. Water Research 6 57 (1972) 126 Ducreux, J., Boulet, R., Petroff, N. and Roussel, J.C. International Journal of Environmental Analytical Chemistry 12 195 (1982) 127 Stottmeister, E., Heudel, P. and Hermenau, H. Chemosphere 17 801 (1988) 128 Gomez-Belinchou, J.T. and Albaiges, J. International Journal of Environmental Analytical Chemistry 30 183 (1987) 129 Mirzayanov, U.S. and Bugrov, Yu.F. Zavod. Lab. 38 656 (1972) 130 Borneff, J. and Kunte, H. Arch. Hyg. Bakt. 153 202 (1968) 131 Jentoft, R.E. and Gouw, T.H. Analytical Chemistry 40 1787 (1968) 132 McKay, J.F. and Latham, D.R. Analytical Chemistry 45 1050 (1973) 133 Vaughan, C.G., Wheals, B.B. and Whitehouse, H.J. Journal of Chromatography 78 203 (1973) 134 Melchiorri, C., Chiacchiarini, L., Grilla, A. and D’Area, S.U.N. Ann. Ig. Mier. 24 279 (1973) 135 Monarcha, S., Conti, R., Scassellati Sforzolini, G. and Savino, A. Iq. Mod. 69 331 (1976) 136 Borneff. J. In Fate of Pollutants in the Air and Water Environments (Ed. Suffet, I.H.), John Wiley, Chichester, Vol. 2, p. 379 (1977) 137 Outkiewicz, T., Ryborz, S. and Maslowski, J. Environ. Prot. Engng 4 263, (1978) 138 Woo, C.S., D’Silva, A.P. and Fassell, D.A. Analytical Chemistry 52 159 (1980) 139 Navra’til, J. D., Sievers, R.E. and Walton, W.F. Analytical Chemistry 49 2260 (1977) 140 Chatot, G., Jequier, W., Jay, M., Fontages, R., and Obaton, P. Journal of Chromatography 45 415 (1969) 141 Frycka, J. J. Chromat. 65 432 (1972) 142 Dunn, B.P. and Stich, H.F. J. Fish. Res. Bd Can. 33 2040 (1976) 143 Harrison, R.M., Perry, R. and Wellings, R.A. Wat. Res. 10 207 (1976) 144 Hellmann, H.Z. Anal. Chem. 275 109 (1975) 145 Acheson, M.A., Harrison, R.M., Perry, R. and Wellings, R. Wat. Res. 10 207 (1976) 146 Caddy, D.E. and Meek, D.M. Technical Report TR36 Water Research Centre, Stevenage Laboratory, Elder Way, Stevenage, Herts (1976) 147 Bowman, M.C. and Benoza, M. Analytical Chemistry 40 535 (1968) 148 Burchfield, H.P., Wheeler, R.J. and Bernos, J.B. Analytical Chemistry 43 1976 (1971) 149 Freed, D.J. and Faulkner, L.R. Analytical Chemistry 44 1194 (1971)
224
Hydrocarbons
150 Robinson, J.W. and Goodbread, J.P. Anal. Chim. Acta 66 239 (1973) 151 Acheson, M.A., Harrison, R.M., Perry, R. and Wellings, R.A. Water Research 10 207 (1976) 152 Hoffmann, D. and Wynder, O. Analytical Chemistry 32 295 (1960) 153 Haenni, E.O., Havard, J.W. and Joe, F.L. J. Ass. Off. Agric. Chem. 45 67 (1962) 154 Natusch, D.F.S. and Tomkins, B.A. Analytical Chemistry 50 1429 (1978) 155 Basu, D.K. and Saxena, J. Journal of Environmental Science and Technology 12 791 (1978) 156 Matsumoto, G. and Hanya. T. Journal of Chromatography 194 199 (1980) 157 Constable, D.J.T., Smith, S.R. and Tanka, J. Environmental Science and Technology 18 975 (1984) 158 Lewis, W.M. Water Treat. Exam. 24 243 (1975) 159 Crathorne, B. and Fielding, M. Pract. Anal. Div. Chem. Soc. 11 155 (1978) 160 Crane, R.I., Crathorne, B. and Fielding N. In Hydrocarbons and Halogenated Hydrocarbons in Aquatic Environments (Environmental Science Research Series, Vol. 16) (Ed. Asghan, B.K.) Plenum, New York (1976) 161 Dong, M., Locke, D.C. and Ferrand, E. Analytical Chemistry 48 368 (1976) 162 Krstulovic, A.M., Rosie, D.M. and Brown, P.R. Analytical Chemistry 48 1383 (1976) 163 Lankmayr, E.P. and Muller, K. Journal of Chromatography 170 139 (1979) 164 Nielson, T.J. Journal of Chromatography 170 147 (1979) 165 Crosby, N.T., Hunt, D.C., Philip, L.A. and Patel, I. Analyst (London) 106 135 (1981) 166 Hagenmaier, H., Feirabend, R. and Jager, W. W. Zeitschrift für Wasser und Abwasser Forschung. 10 99 (1971) 167 Black, J.J., Dymerski, P.P. and Zapisek, W.F. Bull. Environ. Contam. Toxicol. 22 278 (1979) 168 Saxena, J., Basu, D.K. and Kozuchowski, J. Method Development and Monitoring of Polynudear Aromatic Hydrocarbons in Selected US Water, Health Effects Research Laboratory, TR-77-563, No 24, Cincinnati, Ohio (1977) 169 O’Donnell, M. Journal of Environmental Science 1 77 (1980) 170 Perkin–Elmer Ltd. The Analytical Bulletin No. 1 p. 4. (2/2 1981) 171 Standing Committee of Analysts. H.M. Stationery Office, London. Methods for the Examination of Water Associated Materials. Determination of 6 specific polynuclear aromatic hydrocarbons in water (1988) 172 Gilchrist, C.A., Lynes, A., Steel, G. and Whitham, B.T. Analyst (London) 97, 880 (1972) 173 Kay, J.F. and Latham, D.R. Analytical Chemistry 45 1050 (1973) 174 Stepanova, M.I., Ii’ina, R.I. and Shaposhnikov, Zh. Analyt. Khim. 27 1201 (1972) 175 Nowacka-Barezyk, K., Adamiak-Ziemba, J. and Janina, A.Z. Chemia. Anglit. 18 223 (1973) 176 Crane, R.I., Crathorne, B. and Fielding, M. Determination of Polycyclic Aromatic Hydrocarbons in Water, Water Research Centre LR 4407 (1980) 177 Schossner, H., Falkenberg, W. and Althaus, H. Z. Wasser Abwasser Forsch. 16 132 (1983) 178 Scholz, L. and Altmann, H.J. Zeitschrift für Analytische Chemie 240 81 (1968) 179 Schwarz, F.P. and Wasik, S.P. Analytical Chemistry 48 525 (1976) 180 Monarca, S., Causey, B.S. and Kirkbright F.G. Water Research 13 503 (1979) 181 Hood, L.V.S. and Winefordner, J.D. Anal. Chim. Acta 42 199 (1968) 182 Lavalette, D., Muel, B., Habert-Habert, M., René, L. and Latarjet, R. Journal of Chemical Physics 65 2141 (1968)
References
225
183 Gaevaya, T. Ya. and Khesina, A. Ya. Analytical Chemistry USSR 29 1913 (1974) 184 Nurmukamelov, R.N. Russian Chemical Review 53 180 (1969) 185 Kirkbright, G.F. and De Lima, C.G. Analyst (London) 99 338 (1974) 186 Causey, B.S., Kirkbright, G.F. and De Lima, C.G. Analyst (London) 101 367 (1976) 187 Farooq, R. and Kirkbright, G.F. Environ. Sci. Technol. 10 1018 (1976) 188 Sampling and Analysis Procedures for Screening of Industrial Effluents for Priority Pollutants, US Environmental Protection Agency, Environmental Monitoring and Support Laboratory, Cincinnati, Ohio, April (1977) 189 Shabad, L.M. Hyg. Sanit. 36 155 (1971) 190 Blyshak, L.A., Rossi, T.M., Patonen, G., and Warner, I.M. Analytical Chemistry 60 2127 (1988) 191 Baralkiewicz, D., Grameuska, L.T. and Zerbe, S. Gaspodarka Wodna. 48 41 (1988) 192 Afghan, B.K., Wilkinson, R.J., Chow, A., Findley, T.W., Gessler, H.D. and Srikameswaren, K.S. Water Research 18 9 (1984) 193 Hess, G.G., McKenzie, D.E. and Hughes, B.M. Journal of Chromatography 366 197 (1986) 194 Ghaoqui, L. Journal of Chromatography 399 69 (1987) 195 Gurov, F.K. and Novikov, Yu.V. Hyg. Sanit. 36 409 (1971) 196 Hiltabrand, R.T. Marine Pollution Bulletin 9 19 (1978) 197 Payne, J.F. Science 191 945 (1976) 198 Isaaq, H.J., Andrews, A.W., Janini, G.N. and Barr, E.W. Journal of Chromatography 2 319 (1979) 199 International Standards for Drinking Water, third edition, World Health Organization, Geneva, p. 37 (1971) 200 Cathrone, B. and Fielding, M. Proc. Anal. Div. Chem. Soc. 15 155 (1978) 201 Saxena, J., Basu, D.K. and Kozuchowski. J. US National Technical Information Service, Springfield, VA, Report No. PB276635 p. 94 (1997) 202 Saxena, J., Kozuchowski, J. and Basu, D.K. Environmental Science and Technology 11 682 (1977) 203 Benoit, F., Label, G.L. and Williams, D.T. International Journal of Analytical Chemistry 6 277 (1979) 204 McNeil, E.E., Olson, R., Miles, W.F. and Rajalalee, R.J.M. Journal of Chromatography. 132 277 (1977) 205 Grimmer, G., Dettbarn, G. and Schneider, D.W. Wasser Abwasser Forsch. 14 100 (1981) 206 Grimmer, G., Hellman, H., de Jong, B., Kunte, H., Sanneborn, M. and Stroker, I. Zeitschrift für Wasser Abwasser Forsch. 13 108 (1980) 207 Kunte, H. Fresenius Z. Analyt. Chemie 301 287 (1980) 208 Harrison, R.M., Perry, R. and Wellings, R.A. Environ. Sci. Technol. 10 1151 (1976) 209 Weil, L., Grimmer, G., Hellman, H., de Jong, B., Kunte, H., Sanneborn, M. and Stroker, I. Zeitschrift für Wasser und Abwasser Forschung. 13 108 (1980) 210 Sorrell, R.K., Dressman, R.C. and McFarren, E.F. Environment Protection Agency, Cincinnati, Ohio. High Pressure Liquid Chromatography for the Measurement of PAHs in Water, Report No 29344 p. 39 (1977), Paper presented at Water Quality Technology Conference, Kansas City, Missouri, 5/6 Dec. 1977 211 Oyler, A.R., Bodenner, D.L., Welch, K.J., Hukhoxen, R.K., Carlson, R.M., Kopperman, H.L. and Caple, R. Analytical Chemistry 50 837 (1978)
226
Hydrocarbons
212 Hunt. D.C., Wild. P.J. and Crosby. N.T. Water Research 23 643 (1978) 213 Sorrell, R.K. and Reding, R.J. Chromat. 185 655 (1979) 214 Sorrell. R.K., Dressman, R.C. and McFarren, E.F. AWWA Water Quality Technology Conference, Kansas City, MO, December 5–7 1977, American Water Works Association, Denver, Colo, P.3A-3 (1978) 215 Perkin–Elmer Ltd. The Analytical Bulletin No. 1 p. 4 (2/2 1982) 216 Muel, B. and Lacroix, G. Bulletin Chemical Society 2139 (1960) 217 Jager, J. and Kassovitzova, B. Chemy Listy 62 216 (1968) 218 Ogan, K. and Slavin, W.J. Journal of Chromatographic Science 16 517 (1978) 219 International Standards for Drinking Water, third edition, World Health Organization, Geneva, p. 37 (1971) 220 Nelson, A., Aufreet, N. and Readman, J. Anal. Chim, Acta 207 47 (1988) 221 Thieleman, H. Mikrochim. Acta 5 575 (1972) 222 Cannavacinolo. F., Goretti, G., Lagana, A., Petronia, B.B. and Zoccolillo, L. Chromatographia 13 223 (1980) 223 Legana, A., Petronia, B.M. and Rotatori, M. Journal of Chromatography 198 143 (1980) 224 Kadar, R., Nagy, K. and Fremstad, D. Talanta 27 227 (1980) 225 Das, B.S. and Thomas, G.H. Analytical Chemistry 50 , 967 (1978) 226 Khesina, A. Ya. and Petrova, T.B. J. Spectrosc. USSR 18 622 (1973) 227 Andelmann, J.B. and Snodgrass, J.E. CRC Crit. Rev. Environ. Contr. 4 69 (1974) 228 Andelmann, J.B. and Suess, M.J. Bull. Wld Hlth Org. 43 479 (1970) 229 Searl, T.D., King, W.H. and Brown, R.A. Analytical Chemistry 42 954 (1970) 230 Lao, R.C., Thomas, R.S. and Monkman, J.L. Journal of Chromatography 112 681 (1975) 231 Environmental Protection Agency, EPA 610, Environmental Protection Agency USA . (1977) 232 Kasiske, D., Klinkmuller, K.D. and Sonneborn, M.J. J. Chromat. 149 703 (1978) 233 Symons, R.E. and Crick, I. Anal. Chim. Acta 151 237 (1983) 234 Alexander, R., Cumbers, K.M. and Kagi, R.I. Int. J. Environ. Anal. Chem. 12 161 (1982) 235 Stepanova, M.I., Ill’ina, R.I. and Shaposhnikov, Yu. K. Analytical Chemistry USSR 27 1075 (1972) 236 Grzybowski, J., Radecki, A. and Rewkowska, G. Environ. Sci. Technol. 17 44 (1983) 237 Brown, L., Rhead, M.M. and Braven, J. Water Research 18 995 (1984) 238 Brunnock, J.V., Duckworth, D.F. and Stephens, G.G. J. Inst. Petrol. 54 310 (1966) 239 Duewer, D.L., Kowalski, B.R. and Schatski, T.F. Analytical Chemistry 47 1573 (1975) 240 Kawahara, F.K. and Ballinger, D.G. Ind. Eng. Chem. Prod. Res. Dev. 9 553 (1970) 241 Lynch, P.F. and Brown, Ch. W. Environ. Sci. Technol 7 1123 (1973) 242 Mattson, J.S., Mattson, C.S., Spencer, M.J. and Starks, S.A. Analytical Chemistry 49 297 (1977) 243 Erhardt, J.S. and Blumer, M. Environ. Pollut. 3 179 (1972) 244 Clark, H.A. and Jurs, P.C. Analytical Chemistry 47 374 (1975) 245 Rasmussen, D.V. Analytical Chemistry 48 1562 (1976)
References
227
246 Kawahara, F.K. Laboratory Guide for the Identification of Petroleum Products. 1014 Broadway, Cincinnati, Ohio, U.S. Dept of the Interior , Federal Water Pollution Central Administration, Analytical Quality Central Laboratory, 41 pp. (1969) 247 Davies, A.W. Cent. Belge Étud. Docum. Eaux. 330 252 (1971) 248 Caruso, S.C. Devs. Appl. Spectrosc. 6 323 (1967) 249 Peake, E. and Hodgson, G.W. J. Am. Oil Chem. Soc. 43 215 (1966) 250 Peake, E. and Hodgson, G.W. J. Am. Oil Chem. Soc. 44 696 (1967) 251 Adams, I.M. Process Biochem. 2 33 (1967) 252 Institute of Petroleum Standardization Committee. Journal of the Institute of Petroleum (London) 56 107 (1970) 253 Rook, J.J. H O 4 385 (1971) 2 254 McAuliffe, C. Chemical Geology 4 255 (1969) 255 Goma, G. and Durand, G. Water Research 5 545 (1971) 256 Gouw, T.H. Analytical Chemistry 42 1394 (1970) 257 Trent River Authority , Private communication , Trent River Authority , Meadow Lane , Nottingham 258 Ramsdale, S.J. and Wilkinson, R.E. Journal of the Institute of Petroleum (London) 54 326 (1968) 259 Johnson, W. and Kawahara, F.K. Proc. 11th Conf. Great. Lakes Res., p. 550 (1968) 260 Ellerker, R. Wat. Pollut. Control. 67 542 (1968) 261 Dewitt Johnson, W. and Fuller, F.D. Proc. 13th Conf. Great. Lakes Res., p. 128 (1970) 262 Blumer, M. Mar. Biol. 5 195 (1970) 263 Le Pera, M.E. Identification and Characterization of Petroleum Fuels using Temperature-Programmed Gas–Liquid Chromatography, Springfield, VA, National Technical Information Service, Rep. no. AD 646-382 (1966) 264 Duckworth, D.F. Aspects of petroleum pollutant analysis. In Water Pollution in Oil (ed. P. Hepplee), London, Institute of Petroleum, p. 165 (1971) 265 Jeltes, R. Anal. Abstr. 15 3633 (1968) 266 Adlard, E.R. and Matthews, P.H.D. Natur. Physical Science 233 83 (1971) 267 Lur’e, Yu.Yu., Panova, V.A. and Nickolaeva, Z.V. Gidrokhim. Water 55 108 (1971) Ref. Zhur Khim. 199D (1971) (16) Abstract No. 16G258 268 Cole, R.D. Nature (London) 233 546 (1971) 269 Jeltes, R. and Van Tonkelaar, W.A.M. Water Research 6 271 (1972) 270 Jeltes, R. and Van Tonkelaar W.A.M. H O 5 288 (1972) 2 271 Kawahara, F.K. J. Chromatographic Science 10 629 (1972) 272 Lysyj, I. and Newton, P.R. Analytical Chemistry 44 2385 (1972) 273 Lysyj, I. and Newton, P.R. Analytical Abstracts 22 459 (1972) 274 Garra, M.E. and Muth, J. Science and Technology 8 249 (1974) 275 Vos, L., Bridie, A.L. and Herzberg, S. H O 10 277 (1977) 2 276 Crump, G.B. Nature (London) 193 674 (1963) 277 Krieger, H. Gas- u. Wassfach. 104 695 (1963) 278 Lambert, G. Bull. Cent. Belge Stud. Docm. Eaux 20 271 (1966) 279 Matthews, P.J. Water Pollution Control 67 588 (1968) 280 Matthews, P.J. J. Appl. Chem (London) 20 87 (1970) 281 Cross, B.T. and Hart, M. Water Research Association, Medmanham (UK), Internal Laboratory Report 1RL62 (1962) 282 Semenov, A.D., Stradomanskaya, A.G. and Zurbina, L.F. Gidrokhim Mater. 53 51 (1971) Ref. Zhur. Khim 19GD (1971)
228
Hydrocarbons
283 Sauer, W.A. and Fitzgerald, G.E. Environmental Science and Technology 10 893 (1976) 284 Killer, F.C.A. and Amos, R. Journal of Institute of Petroleum (London) 52 315 (1966) 285 Coats, J.P. Journal of Institute of Petroleum (London) 57 209 (1971) 286 Triems, K. Chem. Tech. Berl. 20 596 (1968) 287 Triems, K. and Heinze, O. Anal. Abstr. 16 2556 (1969) 288 Farrington, J.W., Teal, J.M., Quinn, J.G., Wade, T. and Burnes, K. Bulletin of Environmental Contamination and Toxicology 10 129 (1973) 289 Hellman, H. Deutsch Gewaesserk. Mitt. 13 19 (1969) 290 Geyer, D., Martin, P. and Adrian, P. Gas- u. Wasserfach. Wasser Abwasser 119 72 (1978) 291 Peitscher, G.W. Fresenius Z. für Analytische Chemie 327 56 (1987) 292 Freegarde, M. Laboratory Practice 20 35 (1971) 293 Thrusten, A.D. and Knight R.W. Environmental Science and Technology 5 64 (1971) 294 Lloyd. J.B.F. J. Forens. Sci. Soc. 11 83 (1971) 295 Lloyd. J.B.F. J. Forens. Sci. Soc. 11 153 (1971) 296 Lloyd. J.B.F. J. Forens. Sci. Soc. 11 235 (1971) 297 Parker, C.A. and Barnes, W.J. Analyst (London) 85 3 (1960) 298 Bauer, K. and Driescher, H. Fortschr. Wasserchem ihrer Grenzgeb. 10 31 (1968) 299 Danyl, F. and Nietsch, B. Mikrochemie Mikrochem. Acta 39 333 (1952) 300 Nietsch. B. Angew. Chem. 66 571 (1954) 301 Nietsch, B. Gass. Wass. Warme. 10 66 (1956) 302 Nietsch, B. Mikrochim. Acta 171 (1956) 303 Leoy, E.M. Water Research 5 723 (1971) 304 Sinel’nikov, V.E. Kazan. Med. Zh. 3 83 (1968) 305 Environmental Protection Agency Industrial Environmental Research Laboratory, Cincinnati , Ohio, Report No. 32428, p. 256 (1979) 306 John, P., McQuat, E.C. and Sontar, L. Analyst (London) 107 221 (1982) 307 Stuart, R.A. and Branch, R.D. Instrument News (Perkin–Elmer Corporation, Norwalk, Connecticut) 21 No. 2 (1970) 308 Bryan, D.E. and Guinn, V.P. Springfield, Virginia National Information Service, USA. EC Report No GA 9889 , p. 134 (1970) 309 Guinn, V.P. and Bellanca, S.C. Washington USA PO National Bureau of Standards Special Publication No 312, p. 93 (1967) 310 Bryan, D.E., Guinn, V.P. Hacklemann, R.P. and Leukens, H.R. Report Atomic Energy Commission US GA-9889, p. 134 (1970) 311 Lukens, H.R., Bryan, D., Hiatt, N.A. and Schlesinger, H.L. Report Atomic Energy Commission US GULF-RT-A-10684, p. 134 (1971) 312 Lukens, H.R. Report Atomic Energy Commision US GULF-RT-A-10973, p. 74 (1972) 313 Flaherty, J.P. and Eldridge, H.B. Applied Spectroscopy 24 534 (1970) 314 Scott, K.A. and Bean, B.M. US Patent No 3 574 550. Washington Patent Office (1976) 315 Louis, R. Chemie Ingr-Tech. 40 538 (1968) 316 Kubo, H., Bernthal, R. and Wildman, R. Analytical Chemistry 50 899 (1978) 317 Eldridge, H.B. and Flaherty, J.P. Ind. Engng. Chem. Prod. Res. Dev. 9 422 (1970) 318 Budnikov, G.K. and Medyantseva, E.P. Zhur. Analit. Khim. 28 301 (1973) 319 Carsin, J.L. Revue Internationale d’Oceanographic Medicale 48 77 (1977) 320 Hellmann, H.Z. Analytical Chemistry 244 44 (1969)
References
229
321 Braemer, H.C. Proc. Am. Chem. Soc. Div. Wat. Waste Chem. (1966) 322 Gregory, G.R.E.C. and Palmer, P.L. Apparatus for Detecting Oil in Water. Brit. Patent 1 221 066 (London Patent Office) 323 Muskewity, G. Appparatus for the continuous determination of oils in liquid and gaseous media, Ger. Pat. 1 810 604 (Berlin Patent Office) 324 Grabbe, F. Industrieabwasser 45 (1968) 325 Emeschregenossenschaft, Essen. Eff. Wat. Treat. J. 10 667 (1970) 326 Goolsby, A.D. Environ. Sci. Technol. 5 356 (1971) 327 Vasicek, A. Optics of Thin Films, North-Holland Pub. Co., Amsterdam, pp. 26–41 (1960) 328 Mattson, J.S. Environ. Sci. Technol. 5 415 (1971) 329 Fust, H.W., Kreider, R.E. and Gardiner, K.W. Seventeenth Annual ISA Analysis Instrumentation Symposium, Houston, Texas, April (1971). Analysis Instrumentation 9, Publ. Instrument Society of America, Pittsburgh, May (1971), Paper 45 Automated instrumental approach for oil in water. 330 Ahmadijian, M. and Brown, C.W. Environ. Sci. Technol. 7 452 (1973) 331 McMullen, A.I., Monk, J.F. and Stuart, M.J. International Laboratory 60–61 , Jan.–Feb. 5447 (1975) 332 Uchiyama, M. Water Res. 12 299 (1978) 333 Steven, S.F., Smith, J.F., Flego, U. and Renber, J. Water Research 12 447 (1978) 334 Wilson, C.A., Ferreto, E.P. and Coleman, H.J. Am. Chem. Soc. Div. Chem. Pepr. 60 613 (1975) 335 Garza, M.E. and Muth, J. Environmental Science and Technology. 8 249 (1974) 336 Adlard, E.R., Creaser, L.F. and Matthews, P.H. Analytical Chemistry 44 297 (1972) 337 Blumer, M. and Sass, J.J. Science 176 1120 (1972) 338 Boylan, D.B. and Tripp, B.W. Nature (London) 230 44 (1971) 339 McKay, T.R. Proc. Ninth Int. Symp. on Gas Chromatography, Montreux, Switzerland, pp. 33–38 October (1972) 340 Zafiron, O.C., Myers, J. and Freestone, F. Marine Pollution Bulletin 4 87 (1973) 341 Bulten, J.N., Morris, B.F. and Sass, J. J. Bermuda Biological Research, Special Publication No 10. 350 pp . (1973) 342 Zafiron, O.C. and Oliver, C. Analytical Chemistry 45 952 (1973) 343 Hertz, H.S., May, W.E., Chesler, S.N. and Gump, B.H. Environ. Sci. Technol. 10 , 900 (1976) 344 Smith, D.H. Analytical Chemistry 44 536 (1972) 345 Walker, J.D., Colwell, R.P., Hamming, M.C. and Ford, H.I. Bulletin of Environmental Contamination and Toxicology 13 245 (1975) 346 Brown, R.A. and Huffman, H.L. Science 101 847 (1976) 347 Albaigés, J. and Albrecht, P. International Environmental Analytical Chemistry 6 171 (1979) 348 Bunnock, J.V., Duckworth, D.F. and Stephens, G.G. Inst. Petrol. 54 310 (1968) 349 Bailey, N.J.L., Jobson, A.M. and Rogers, M.A. Chemical Geology 11 203 (1973) 350 Henderson, W., Wollrab, V. and Eglington, G. Advances in Organic Geochemistry 1968 ( eds Schenk, P.A. and Havenaar, I.), p. 181, Pergamon, Oxford (1969) 351 Gallegos, E.J. Anal. Chem. 43 1151 (1971) 352 Rubinstein, I., Strausz, O.P., Spyckerelle, C, Crawford, R.J. and Westlake, D.W.S. Geochem. Cosmochim. Acta 41 1341 (1977) 353 Albaigés, J., Borbon, J. and Salgre, P. Tetrahedron Lett. 595, (1978) 354 Rubinstein, I. and Albrecht, P. J. Chem. Soc. Chem. Commun. 957 (1975)
230
Hydrocarbons
355 Van Dorsselaar, A., Albrecht, P. and Ourisson, G. Bull. Soc. Chim. France 165, (1977) 356 Kimble, B.J., Maxwell, J.R., Philip, R.P., Eglington, G., Albrecht, P., Ensminger, A., Arpino, P. and Curisson, G. Geochim. Cosmochim. Acta 38 1165 (1974) 357 Schuldiner, J.A. Analytical Chemistry 23 1676 (1951) 358 Herd, M. Analyst (London) 78 383 (1953) 359 Done, J.M. and Reid, W.K. Separation Science 5 825 (1970) 360 Mattson, J.S. and Mark, H.B. Analytical Chemistry 42 234 (1970) 361 Mattson, J.S. and Mark, H.B. Environ. Sci. Technol. 3 161 (1969) 362 Mark, H.B., Yu, T.C., Mattson, J.S. and Kolpack, R.L. Environ. Sci. Technol. 6 833 (1972) 363 Baier, R.E. J. Geophys Res.. 77 5062 (1972) 364 Mattson, J.S. Analytical Chemistry 43 1872 (1972) 365 Ahmed, S.M., Beasley, M.D., Etomson, W.C. and Hites, R.A. Analytical Chemistry 46 1858 (1974) 366 Ahmadjian, M and Brown, C.V. Analytical Chemistry 48 1257 (1976) 367 Zitko, V. and Carson, W.V. Tech. Report Fish. Res. Bd. Can. No. 217, 29 pp . (1970) 368 Michalik, P.A. and Gordon, D.C. Tech. Report Fish. Res. Bd. Can. No 284, 26 pp . (1971) 369 Levy, E.M. Water Research 5 723 (1971) 370 Levy, E.M. Water Air Soil Pollution 1 144 (1972) 371 Levy, E.M. and Walton, A. J. Fish. Res. Bd . Can. 30 261 (1973) 372 Hargrave, B.T. and Phillips, G.A. Environmental Pollution 8 193 (1975) 373 Freegarde, M., Hatchard, C.G. and Parker, C.A. Laboratory Practice 20 35 (1971) 374 Lieberman, M. Report U.S. Environmental Protection Agency, EPA-RZ73102, 184 pp. (1973) 375 Millard, J.P. and Arvesen, J.C. Applied Optics 11 102 (1972) 376 Pierre, L.J. Applied Optics 12 2035 (1973) 377 Martin, P. and Geyer, D., Korrespond. Abwasser 21 202 (1974) 378 Dadeshev, Kh.K. and Agamirova, S.I. Vop. Issled. Neft. I nefteprod. razabotki protseeor pererabotki neft. Obsled. Zav. Stanovok No. 1, p. 81 (1957) 379 Rychkova, V.I. Elect. Sta. Mosk. 40 76 (1969) 380 Alekseeva, V. and Gol’dina, Ts.A. Zav. Lab. 16 35 (1950) 381 Yudilevich, M.M. Luminescence Methods for the Determination of the Content of Mineral Oils in Water (in Russian), Gosudarst. Energet. Izdatel, Moscow (1959) 382 Yudilevich, M.M. Luminescence Method and Apparatus for Analysis of Water– Oil Emulsions (in Russian), Metody Lyumineestsentn. Analiza (Minsk: Akad. Nuak Belorum S.S.R.) Sbornik, 87 (1960) 383 Yudilevich, M.M. In Russian, Peredoye Metody Khim. Teknol. i Kontrolya Proizv. 280 (1964) 384 Yudilevich, M.M. In Russian, Vodopodgotovka, Vod. Rezhim Khim Kontrol. Parosilovykh Ustanovkakh, S.B. Statei, No. 2, 173 (1966) 385 Kyrge, Kh. In Russian, Morsk. Flot. 19 31 (1959) 386 Leger, A. Fr. Pat. 1 560 844 (Paris, Patent Office) 387 Leonchenkova, E.T. In Russian, Obogasshch. Rud. 5 24 (1960) 388 Shklyar, I.V. In Russian, Trudy vses. Nauchno-issled, geologora zved. Inst. No. 155, 341 (1960) 389 Pochkin, Yu. N. and Massino, O.A. In Russian, Kazan. Ed. Zh. 3 , 81 (1968)
References
390 391 392 393 394 395 396 397 398 399
400 401 402 403 404 405 406 407 408 409 410 411 412 413 414
231
Millson, M.F. J. Chromatography, 50 , 155 (1970) Taras, M.J. and Blum, K.A. Water Pollution Control 40 404 (1968) Maxcy, R.B. J. Water Pollution. Contr. Fed. 48 2809 (1976) Cook, P.P., Duvall, P.M. and Bourke, R.C. Water and Sewage Works. Ref. No. 1978 R71-R72 and R74-R78 (1978) US Environmental Protection Agency. Methods for Chemical Analysis of Water and Wastes, EPA 625 (6.74 003 STORET NO. 00556) (1974) Pomeroy, R. and Wakeman, C.M. Ind. Engng. Chem. Anal. Ed. 13 795 (1941) Gilcreas, F.W. Sewage Ind. Wastes 25 1379 (1958) Ullman, W.W. and Sanderson, W.W. Sewage Ind. Wastes 31 8 (1959) Cleverley, B.J. Forens. Sci. Soc. 8 69 (1959) Standing Committeee of Analysts. H.M. Stationery Office, London. Methods for the Examination of Waters and Associated Materials 1984, 19 pp . (P22B : C ENV) (1984). The determination of hydrocarbon oils in waters by solvent extraction. Infra-red absorption and gravimetry (1983) National Water Council, London. Methods for the Examination of Waters and Associated Materials 1988. Determination of oils and fats in wastewater by filtration solvent extraction and gravimetry (1987) Louch, D., Motlagh, S. and Pawliszyn, J.J. Anal. Chem. 64 1187 (1992) Okahata, Y. and Ebato, H. Anal. Chem. 61 2185 (1989) Wohltjen, H. Sens. Actuators 5 307 (1984) Krska, R., Kellner, R., Schiessl, U., Tacke, M. and Karzir, A. Appl. Phys. Letters 63 1868 (1993) Krska, R., Rossenberg, E., Taga, D., Kellner, R., Messicar, A. and Katzir, A. Applied Phys. Lett. 61 1778 (1992) Kvasnik, F. and McGrath, A.D. SPIE Chem. Biochem. Environ. Sensors 1172 75 (1989) DeGrandpre, M.D. and Burgess, L.W. Appl Spectros. 44 273 (1990) DeGrandpre, M.D. and Burgess, L.W. ISA Trans. 28 71 (1988) Burck, J., Conzen, J.P., Bechaus, B. and Ache, H.J. Sens. Actuators B. 18–19 291 (1994) DeGrandpre, M.D. and Burgess, L.W. Anal. Chem. 60 2582 (1988) Burck, J., Conzen, J.P. and Ache, H.J. Fresenius Z. Anal. Chem. 342 394 (1992) Burck, J., Conzen, J.P. and Ache, H.J. Appl. Spectrosc. 47 753 (1993) Kusada, K., Shiraki, K. and Miva, T. Anal Chim. Acta 224 1 (1989) Symons, R.K. and Crick, I. Anal. Chim. Acta 151 237 (1983)
Chapter 3
Oxygen-containing compounds
3.1 Carboxylic acids 3.1.1 Natural (non-saline) waters There are two sources of fatty acids in river water, man-made contamination and naturally occurring. Regarding the latter the determination of free and bound fatty acids present in aquatic systems is important first because fatty acids are sufficiently diverse in structure that they can be used to determine the source and cycling of organic carbon [1–5] and second because fatty acids in vivo function primarily as structural compounds of membranes and energy storage products. In the reduced state the amount and type of fatty acids may be indicative of the trophic status of the ecosystem at the time of fatty acid formation [2, 6]. Unsaturated, short-chain (up to C ) and microbial fatty acids are 20 indicative of productive systems, whereas long-chain acids dominate in oligotrophic systems. Fatty acids have been shown to constitute 5–10% of the weight of humic/fulvic acid structure [7]. As an integral portion of the structure of these refractory materials, fatty acids can be used to determine the source of organic carbon and the physical/chemical characteristics of these materials. The behaviour of humic and fulvic acids may determine the transport of toxic trace metals and anthropogenic organics. 3.1.1.1 Spectrophotometric methods Lactic acid Stradomskaya and Goncharova [8] describe a Spectrophotometric method for the determination of lactic acid in amounts down to 5µg in natural water. The method is based on oxidation of lactic acid to
Carboxylic acids
233
acetaldehyde and reaction of the latter with 1-naphthol to form a coloured product. Amino acids, aldehydes (other the acetaldehyde), organic acids and inorganic ions do not interfere. Benzoic acid Goncharova and Khomenkô [9] determined benzoic acid in ether extracts of natural and contaminated waters. The benzoic acid was back-extracted from the ether into 0.1M aqueous sodium hydrochloric acid and evaluated spectrophotometrically at 230nm. Beer’s law is obeyed with 200–140µg of benzoic acid in 20mL. Phenol, benzene and carboxylic acids (succinic, citric, lactic, oxalic, malic, glutaric, propionic, acetic and formic) do not interfere. The presence of humic and fulvic acids has little effect since they are not extracted into ether. Malic acid Smotrakov and Stradomskaya [10, 11] determined this acid spectrophotometrically in natural water. Malic acid is oxidized to carbonyl compounds with potassium permanganate in sulphuric acid medium and then the carbonyl compounds are reacted with 2,4dinitrophenylhydrazine. The 2,4-dinitrophenylhydrazones are then dissolved in potassium hydroxide in 50% ethyl alcohol and evaluated at 554nm. Phenylglyoxylic and mandelic acids Hatsue and Masayuki [12] determined these acids spectrophotometrically as follows. The sample (0.5mL) is acidified with 0.05mL of 1M hydrochloric acid and shaken with 5mL of ethyl ether for 10min. A portion of the ether layer is evaporated to dryness on a water bath at 70°C. To the residue is added 4mL of concentrated sulphuric acid–40% aqueous formaldehyde (100:1) reagent, and after 15–60min the extinctions at 350 and 450nm are measured. The extinction due to phenylglyoxylic acid has a maximum at 350nm and that due to mandelic acid a maximum at 450nm, and the extinctions are proportional to the amounts of each acid and are additive when mixtures of the acids are analysed. Methacrylic acid Krotova [13] determined this acid spectrophotometrically. A portion of the effluent was extracted with toluene or benzene to remove N-phenyl2-naphthylamine, µa-dimethylbenzyl alcohol and acetophenone and a 50mL sample was adjusted to pH8 (to prevent polymerization) and
234
Oxygen-containing compounds
subjected to distillation in a stream of nitrogen to remove benzene, toluene, butadiene, acetone, or other volatile substances. The solution was diluted to 50mL with water and adjusted to pH4–5 with 0.1M hydrochloric acid. The extinction is measured at 208nm. Formic acid, formaldehyde Jordan [14] has described a procedure for the determination of traces of formic acid and formaldehyde in river water. Formic acid plus formaldehyde are determined after reducing the acid with nascent hydrogen: formaldehyde alone is determined without reduction. The chromatographic acid spectrophotometric method is carried out in 6– 7.5M sulphuric acid. The limit of determination is 50µg L-1 without preconcentration and 50ng L-1 with preconcentration by extraction with diethyl ether. 3.1.1.2 Gas chromatography Kawahara [15, 16] carried out the microdetermination of pentafluorobenzylester derivatives of benzoic, oleic, linoleic and undecanoic acids by electron-capture gas chromatography. The method was applied to chloroform extracts on natural water. Khomenkô and Goncharova [17] separated and concentrated the following acids dissolved in natural water: oxalic, malonic, succinic, glutaric, adipic, fumaric, aconitic, lactic, malic, tartaric, citric, pyruvic and gallic. Ethyl ether–butanol (1:1) and isobutyl alcohol and ethyl acetate were used as solvents. The preferred method was extraction (3 × 5min) at pH2 into an equal volume of butanol. Van Huyssteen [18] completely separated normal and iso acids on glass columns (2.13m × 3mm i.d.) packed with 3% of FFAP (a reaction product of polyethylene glycol 20 000 and 2-nitrophthalic acid developed by Varian Aerograph) on Chromosorb 101 (80–100 mesh) at 180°C in an instrument equipped with a dual flame ionization detector; nitrogen was used as carrier gas (77mL min-1). Bethge and Lindstroem [19] first removed metal cations from a 10mL sample of water by elution with water from a Dowex 50W-X8 ionexchange column and the eluate was titrated to pH8 with standard tetrabutylammonium hydroxide. A calculated amount (as determined from the titration) of hexanoic acid was added as internal standard, the solution was concentrated to a syrup, the syrup was dissolved in acetone, and a-bromotoluene was added in slight excess. After 2h to ensure complete reaction, 1µL of the acetone solution was injected into a stainless steel column (2m × 2mm) packed with 3% of butane-1,4-diol succinate with nitrogen (30mL min-1), as carrier gas and flame ionization
Carboxylic acids
235
detection was used. The column was kept at 120°C for 17min, then temperature programmed to 150°C at 2.5°C min-1. Down to 50µM concentration of the acids could be determined by this procedure. Richard and Fritz [20] employed macroreticular XAD-4 resin aminated with trimethylamine for the concentration, isolation and determination of acidic material from aqueous solutions. Acidic material is separated from other organic material by passing the water sample through a resin column in hydroxide form; other organic compounds are removed with methanol and diethyl ether. The acids are eluted with diethyl ether saturated with hydrogen chloride gas. After concentration the eluate is treated with diazomethane and the esters formed are separated by gas chromatography. Richard et al. [21] concentrated and determined organic acids in river water samples. Concentration was achieved by anion-exchange chromatography with subsequent elution, esterification to methyl derivatives and gas chromatography. The process described was shown to work well for a variety of complex aqueous samples and is simple and reasonably quick. Vairavamurthy et al. [22] determined acrylic acid by electron-capture capillary gas chromatography after extraction with tri-ni-phosphine oxide and derivatization with pentafluorobenzyl bromide. Mebran et al. [23] determined acetic acid by direct aqueous injection gas chromatography. Fatoki and Vernon [24] used capillary gas chromatography of chloroform extracts of natural water to determine free fatty acids. Olsen et al. [25] used a combination of isotope dilution gas chromatography and Fourier transform infrared spectroscopy to analyse methyl esters of several carboxylic acids including malonic, hexabenzene carboxylic, octanoic and octanedioic acids. 3.1.1.3 Thin-layer chromatography Khomenkô et al. [26, 27] used thin-layer chromatography for determining non-volatile organic acids dissolved in natural water. The organic acids are extracted from the water and concentrated, then separated on a silica gel column into four groups which are concentrated to 0.1–0.2mL and thin-layer chromatography is carried out on layers of silica gel KSK previously air-dried for 20min and activated for 30min at 105°C. The acids in the first, second and third fractions are developed in butanol– benzene–acetic acid (10:20:3, 15:85:2 and 15:35:8 respectively) and in the fourth fraction in ethyl acetate–water–formic acid (9:1:1). After drying the chromatograms for 1.5h at 120°C, the organic acids are detected by spraying with 0.4% solution of bromocresol green in 20% ethanolic alkali and the spot areas are measured for a semiquantitative determination of the acids.
236
Oxygen-containing compounds
3.1.1.4 High-performance liquid chromatography Application of high-performance liquid chromatography to the resolution of complex mixtures of fatty acids in water [28, 29] has provided an alternative to the high-temperature separation obtained by gas chromatography. Both techniques have similar limits of detection, but lack the ability to analyse environmental samples directly. Analysis requires that the fatty acids be separated from the organic and inorganic matrices, followed by concentration. Typically, these processes can be accomplished simultaneously by the appropriate choice of methods. Initial isolation of the fatty acids is based on the relative solubility of the material of interest in an organic phase compared to the aqueous phase. Secondary separation is determined by the functional group content and affinity for a solid support. Brown [30] has described a technique for the routine determination of acrylic acid monomer in natural and polluted waters which uses high-performance liquid chromatography for separation for interferences and ultraviolet detection for quantification. Hullett and Eisenreich [31] used high-performance liquid chromatography for the determination of free and bound fatty acids in river water samples. The technique involves sequential liquid– liquid extraction of the water sample by 0.1M hydrochloric acid, benzene–methanol (7:3) and hexane–ether (1:1). The resultant extract was concentrated and the fatty acids were separated as a class on Florasil using an ether– methanol 1:1 and 1:3 elution. Final determination of individual fatty acids was accomplished by forming the chromatophoric phenacyl ester and separating by highperformance liquid chromatography. Bound fatty acids were released by base saponification or acid hydrolysis of a water sample from which the fatty acids had been removed by solvent extraction. These workers [31] defined free fatty acids as those acids which are readily extractable into organic solvents without sample pretreatment with strong acid or base. Bound fatty acids are those acids associated with mineral and humic/fulvic acid surfaces, esterified with mineral and humic/fulvic acid surfaces esterified to humic/fulvic acids, or entrapped in natural macromolecules and released with strong acid or base. Table 3.1 gives the net retention times and relative retention of analytical standards used to characterize the high-performance liquid chromatographic behaviour. The values indicate the baseline resolution is obtained from a homologous series of fatty acids. However, when a complex mixture containing branched and
Carboxylic acids
237
Table 3.1 Retention index for fatty acid separation by HPLC and GLCa,b
Source: Reprinted with permission from Hullett and Eisenreich [31]. Copyright (1979) American Chemical Society. Notes a All time values are given in seconds. b Relative index is calculated relative to C14.0. c Two µBondapak C18 columns in series. d Single µBondapak C16 column.
unsaturated fatty acids was chromatographed, identification and quantification became more difficult. The addition of methyl branching decreased the retention relative to the straight chain saturated homologue. Alkyl substitutions intro duce dipoles which react with the permanent dipole moments of the mobile phase, allowing greater solubility of the solute. Unsaturation decreases the retention of the solute by allowing specific interactions to occur between the polar mobile phase and the π electrons. This increases the solubility of the fatty acids in the mobile phase while decreasing
238
Oxygen-containing compounds
the solubility in the non-polar stationary phase, resulting in a net decrease in retention of the solute by the stationary phase. The good resolution of unsaturated and branched-chain fatty acids obtained by high-performance liquid chromatography is important in the determination of the source of organic carbon. These fatty acids were separated by high-performance liquid chromatography. Figure 3.1 shows high-performance liquid chromatograms obtained for the two Florasil fractions (ether–methanol (1:1) and (1:3)) of a river water sample analysed for free fatty acids. The chromatograms show good separation of the fatty acid material from other compounds. The major fatty acids determined by high-performance liquid chromatography were C >C >C >C >C , while the major fatty 16.0 18.2 14.0 17.0 18.0 acids determined by liquid chromatography were C >C >C >C 16.0 16.1 18.0 17.0. Nimura et al. [32] used 1-pyrenyl diazomethane as a labelling reagent for the liquid chromatographic determination of down to 20–30fmol carboxylic acids including palmitate, 1-pyrenyl palmitate, lactate, propionate and formate. The fluorescent reagent is excited at 340nm and emits at 395nm. Kieber et al. [276] determined formate in natural waters including sea water by a procedure based on coupled enzymatic high-performance liquid chromatography. The procedure is capable of determining down to 4–10µm carboxylic acids. In this procedure, formate is oxidized with formate dehydrogenase and this is accompanied by a corresponding reduction of ßnicotinamideadenine dinucleotide (ß(NAD) +) to reduced ßNADH. ßNADH is quantified by high-performance liquid chromatography. Kishida et al. [33] carried out a specific determination of ascorbate. The ascorbate is derivitivized to dehydroascorbic acid bis(dinitrophenyl) hydrazone (osazone) using 2,4-dinitrophenyl hydrazine as the reagent. The osazone is then determined by high-performance liquid chromatography. 3.1.1.5 Ion-exclusion chromatography Okada [34] used Redox suppressor for the ion-exclusion chromatography of carboxylates using a conductiometric detector on the column. 3.1.1.6 Ion chromatography Saari Nordhaus and Anderson [35] studied the chromatographic separation on a mixed-mode stationary phase of mixtures of carboxylic acids and inorganic anions and showed that by careful selection of eluent pH and ionic strength simultaneous determinations could be carried out
Carboxylic acids
239
Fig. 3.1 HPLC chromatograms of the ether–methanol (1:1) (top) and (1:3) (bottom) fractions. Flow rate 1.0mL min-1; attenuation 0.05AUFS. Peak identification: (1) phenacyl bromide; (2) iC12.0; (3) C12.0; (4) aC13.0; (5) C20.4; (6) C14.0; (7) C18.2; (8) aC15.0; (10) iC16.0; (11) C16.0; (12) aC17.0; (13) C17.0; (14) C18.0. (Reprinted with permission from Hullett and Eisenreich [31]. Copyright (1979). American Chemical Society.)
240
Oxygen-containing compounds
of acetic, lactic, propionic, butyric and isobutyric acids, and also of chloride, bromide nitrile, nitrate, phosphate, selenite and sulphate. Hirajama and Kuwamoto [36] carried out a numerical analysis of the elution behaviour of substituted benzoate anions in ion chromatography. Anions studied included benzoic, chlorobenzoic, various anisates, dimethylbenzoic, salycylic, hydroxybenzoic, resorcylic, protocatechuic and gallic acids. Berglund et al. [37] used two-dimensional conductiometric detection with sequential suppressed and single-column detection in the ion chromatography of glyoxylate and malonate, also fluoride, borate, chloride, arsenate and nitrate in natural waters. 3.1.1.7 Fluorescence spectrometry Bustin and West [38] determined traces of citrate in aqueous systems. The citrate is converted into ammonium 2,6-dihydroxyisonicotinate and the resulting blue fluorescence gives a measure of the amount of citrate. The calibration graph is rectilinear for 0.01–10µg citrate mL-1, the optimum working range is 0.1–10µg mL-1. There was no interference from species normally encountered in water samples. 3.1.1.8 Potentiometric methods Hasson et al. [39] used a potentiometric gas sensing probe for the selective determination of acetate ion in natural waters. 3.1.1.9 Differential pulse polarography Kopanica et al. [40] have described a polarographic determination of small quantities of lactic acid and pyruvic acid (a-ketopropionic acid) in polluted waters. For differential pulse polarographic determination of pyruvic acid, a damping solution of pH4–5 is used with the potential of the polarography wave of -1.10V and a peak width of 110mV. Determination of lactic acid is similarly performed after oxidation by potassium permanganate to pyruvic acid. The method has high precision. 3.1.1.10 Miscellaneous Other procedures for determining fatty acids in water, sediments and biota involve liquid–liquid extraction, liquid–solid adsorption chromatography followed by gas–liquid chromatographic analysis [41– 43]. Liquid extractions have been performed with methanol–chloroform [44], methylene chloride [45] and benzene methanol [46, 47]. Typical
Carboxylic acids
241
liquid–solid adsorbents are silicic acid. Standard gas chromatographic separations for complex mixtures employ non-polar columns packed with OV-1, OV-17, OV-101 or SE-30, or glass capillary columns containing similar phases. 3.1.1.11 Preconcentration Richard and Fritz [20] employed macroreticular XAD-4 resin aminated with trimethylamine for the concentration, isolation and determination of acidic material from aqueous solutions. Acidic material is separated from other organic material by passing the water sample through a resin column in hydroxide form; other organic compounds are removed with methanol and diethyl ether. The acids are eluted with diethyl ether saturated with hydrogen chloride gas. After concentration the eluate is treated with diazomethane and the esters formed are separated by gas chromatography. 3.1.2 Rain water
3.1.2.1 Gas chromatography Kawamura and Kaplan [48] have described a sensitive method for measuring volatile acids (C –C ) in rain and fog samples using ?1 7 bromophenacyl esters and a high-resolution capillary gas chromatograph employing fused silica columns. Experiments showed that the measured concentrations of volatile acids in spiked rain samples increased linearly in proportion to the concentrations of volatile acids added. Relative standard deviations were less than or equal to 18% for C , C and C acids. 1 2 3 The distributions of volatile acids in Los Angeles rain and fog samples are discussed. 3.1.2.2 High-performance liquid chromatography The coupled enzymatic high-performance liquid chromatographic method [276] referred to in Section 3.1.1.4 has been used to determine down to 4–10µM of formate in rain. 3.1.2.3 Ion-exclusion chromatography Backman and Peden [49] used ion-exclusion chromatography to determine weak carboxylic acids in rain water. Citrate, formate and acetate were identified.
242
Oxygen-containing compounds
3.1.3 Sea water 3.1.3.1 Gas chromatography Quinn and Meyers [50] discuss a gas–liquid chromatographic method for the determination of dissolved organic acids in sea water. 3.1.3.2 Atomic absorption spectrometry Treguer et al. [51] determined total dissolved free fatty acids in sea water. The sample (1 L) was shaken with chloroform (2 × 20mL) to remove the free fatty acids and the extract evaporated to dryness under reduced pressure at 50°C. Chloroform–heptane (29:21) (2mL) and fresh copper reagent (M triethanolamine–M acetic acetate–6.8% CuSO ·5H O) solution (9:1:10) (0.5mL) was added to the residue. The 4 2 solution was shaken vigorously for 3min and centrifuged at 3000rpm for 5min. A portion (1.6mL) of the organic phase was evaporated to dryness and 1% ammonium diethyldithiocarbamate solution in isobutylmethyl ketone (2mL) was added to the residue to form a yellow copper complex. The copper in the solution was determined by atomic absorption spectrophotometry at 324.8nm (air–acetylene flame). Palmitic acid was used to prepare a calibration graph. The standard deviation for samples containing 30µg L-1 of free fatty acids (as palmitic acid) was ± 1µg L-1. 3.1.3.3 High-performance liquid chromatography Goncharova and Khomenkô [52] have described a column chromatographic method for the determination of acetic, propionic and butyric acids in sea water and thin-layer chromatographic methods for determining lactic, aconitic, malonic, oxalic, tartaric, citric and malic acids. The pH of the sample is adjusted to 8–9 with sodium hydroxide solution. It is then evaporated almost to dryness at 50–60°C and the residue washed on a filter paper with water acidified with hydrochloric acid. The pH of the resulting solution is adjusted to 2–3 with hydrochloric acid (1:1), the organic acids are extracted into butanol, then back-extracted into sodium hydroxide solution; this solution is concentrated to 0.5–0.7mL, acidified, and the acids separated on a chromatographic column. The coupled enzymatic high-performance liquid chromatographic method [276] referred to in Section 3.1.1.4 has been used to determine down to 0.1–0.8µM of formate in sea water.
Carboxylic acids
243
3.1.3.4 Dif fusion method Xiao et al. [53] determined nanomolar quantities of individual lowmolecular-weight carboxylic acids (and amines) in sea water. This method is based on the diffusion of acids across a hydrophobic membrane to concentrate them and separate them from inorganic salts and most other dissolved organic compounds. Acetic, propionic, butyric, valeric, pyruvic, acrylic and benzoic acids were all found in measurable amounts in sea water. 3.1.4 Waste waters 3.1.4.1 Gas chromatography Gas chromatography has been used for the determination of acetic acid in industrial waste waters [54], and short chain C –C fatty acids in 1 4 anaerobic digester samples. Pilipenko et al. [55] quantitatively determined carboxylic acids in waste waters by gas chromatograhy of their methyl esters. 3.1.4.2 Miscellaneous Narkis and Henfield-Furie [56] have described a procedure for the determination of volatile organic acids in raw municipal waste water. Dzidic et al. [57] used fluoride ion chemical ionization mass spectrometry to determine naphthenic acids in refinery waste waters. 3.1.5 Trade effluents 3.1.5.1 Spectrophotometric methods Abietic type resin acids have been determined in hexane extracts of kraft mill effluents [58]. The effluent was adjusted to pH2.4–2.7 prior to Spectrophotometric evaluation at 490nm of the levoprimaric acid1,amino-2-naphthol-4-sulphonic acid–potassium ferricyanide reaction product. Down to 1mg L-1 of abietic resin acids could be determined in the water sample. Rapid methods of determining formic acid and formaldehyde have been described [14] in which the acid is reduced with nascent hydrogen to formaldehyde. The formaldehyde is determined by the chromotropic acid method in 6–7.5M sulphuric acid. A detection limit of 0.05µg L-1 can be achieved without preconcentration. Preconcentration by extraction with diethyl ether decreases the lower detection limit to 0.05ng L-1. The
244
Oxygen-containing compounds
method has been successfully used for a number of effluent water samples. 3.1.5.2 Gas chromatography Gas chromatography has been used for the determination of acetic acid in industrial waste waters [54], and short-chain C –C fatty acids in 1 4 anaerobic digester samples [18] and dilute aqueous solutions. Van Huyssteen [18] completely separated normal and iso acids on glass columns (2.13m × 3mm i.d.) packed with 3% FFAP on Chromosorb 101 (80–100 mesh) at 180°C in an instrument equipped with a dual-flame ionization detector; nitrogen was used as carrier gas (77mL min-1). C –C fatty acids have been determined in wood and paper 1 4 processing effluents as ?-bromophenacyl esters using glass capillary chromatography and electron-capture detection [59]. A comparison was made of three techniques to treat the samples. Although there was a high response of the electron-capture detector to the ?-bromophenacyl esters of the C –C carboxylic acids, this technique could not be used to its full 1 4 potential. Electron capture was, however, preferred to flame ionization detection. In terms of application in pulp bleaching, to characterize degradation products from wood carbohydrates, formic acid and acetic acid have to be determined; extractive alkylation of tetrahexylammonium ion pairs at pH7, when used for analysis of a kraft paper sample, showed levels of 54 and 11mg L-1 respectively of these acids. Voss and Rapsomatiotis [60] determined resin and fatty acids in paper pulp mill effluents by a solvent extraction–gas chromatographic method. Resin and fatty acids were isolated from the effluent sample by solvent extraction with methyl tert-butyl ether at pH9 and derivatized with diazomethane. The methylated extract was analysed by split capillary chromatography with flame ionization detection. The detection limit was 5ug L-1. The method was applied to fatty and resin acid determination in combined pulp mill effluent entering and leaving the aerated lagoon treatment system of a softwood bleached kraft pulp mill. 3.1.5.3 Miscellaneous McKaveney and Byrnes [61] have described an apparatus incorporating semiconductor electrodes (SCEs) which they have used for the measurement of the concentration of organic acids in pickling bath effluents. Genkin and Zel’manova [62] determined volatile fatty acids and their salts in effluents from aniline dye manufacture. To determine acids the sample was mixed with an equal volume of acetone and titrated potentiometrically (glass electrode and SCE). If mineral acids
Carboxylic acids
245
are absent the inflection in the curve corresponds to the organic acids, but if mineral acids are present this inflection is preceded by one corresponding to these acids. To determine the salts of organic acids, the acids are freed by addition of hydrochloric acid and the solution is titrated similarly. 3.1.6 Sewage effluents 3.1.6.1 Gas chromatography Gas chromatography is a very attractive possibility for volatile acids determination, since it makes the separation of the individual acids for qualitative and quantitative determination in situ possible. In practice, many difficulties in analysing volatile acids in aqueous systems, resulting mainly from the presence of water, have been reported [18]. The volatile acids’ high polarity as well as their tendency to associate and to be adsorbed firmly on the column require esterification prior to gas chromatography determination. The presence of water interferes in esterification so that complex drying techniques and isolation of the acids by extraction, liquid–solid chromatography, distillation and even ion exchangers have to be used [63–66]. The introduction of the more sensitive hydrogen flame ionization detector has made possible the analysis of dilute aqueous solutions of organic acids by gas–liquid chromatography. Problems, such as ‘ghosting’ at high acid concentrations and an excessive tailing effect of the water in dilute solutions, masking the components, have been reported for aqueous solutions [67]. Subsequently phosphoric [68] or metaphosphoric acids [69] were added to the liquid phase, resulting in more reproducible column performance and reduced ‘ghosting’. Addition of formic acid to the carrier gas was recommended by Cochrane [70] to overcome all the problems normally associated with analysing free fatty acids by gas chromatography. Baker [71] used an FFAP column for direct injection of dilute aqueous solutions of acids. The acetic acid peak was not clear and the ability of this column to separate normal and iso fatty acids was not reported. Van Huyssteen [18] successfully used a Chromosorb 101 column coated with 3% FFAP for separation of volatile acids by direct injection of synthetic aqueous solutions and anaerobic digester samples, which were first centrifuged and acidified to pH1–2 with hydrochloric acid. His column affected complete separation of the C – 2 C straight and branched short-chain fatty acids from synthetic 6 aqueous solutions, but less sharpened peaks were obtained from anaerobic digester samples. The response with acetic acid
246
Oxygen-containing compounds
approximated that of the other acids; additional peaks, probably alcohols, appeared between the acid peaks. To eliminate ‘ghosting’ 1–2 µL water was injected between samples. Hindin [72] made sewage samples alkaline with sodium hydroxide prior to evaporation of water for at least 10-fold concentration; 20% metaphosphoric acid was then added and the samples centrifuged. Hindin [72] used a Carbowax 20M plus phosphoric acid on Chromosorb W column. 0.5µg L-1). Dressman et al. [96] are of the opinion that purge and trap methods are very amenable to gas chromatographic–mass spectrometric confirmation of the identity of volatile compounds and at levels lower than can be obtained with liquid–liquid extraction methods. The lower minimum detectable concentrations are attainable because virtually all of the trihalomethanes purged from the sample are transferred to the gas chromatographic column and the detector – without the relatively nonvolatile interferences coextracted by liquid–liquid extraction methods and without introducing solvent-related interference.
388
Halogen-containing compounds
The liquid–liquid extraction methods rarely provide a compound sufficiently concentrated for gas chromatography–mass spectrometry and it may be necessary to resort to a solvent enrichment technique. Enriching the solvent by evaporation commonly results in interference from concentrated solvent impurities to the extent that mass spectral analysis cannot be performed, or in loss of the more volatile components such that actual enrichment is not achieved. Nicolson et al. [132, 139] have described a convenient easily automated method for the analysis of haloforms and some other volatile organohalides in potable water. This direct aqueous injection method has a detection limit at or below 1µg L-1 for haloforms. Simultaneous analysis of finished water samples with direct aqueous injection and the gas sparging method revealed hitherto unknown aspects of water treatment chemistry. While the gas sparging technique measures only the free haloforms present in the potable water, they showed that the direct aqueous injection method quantitates the total potential haloforms that can form after chlorination. Comparison of determinations of haloforms, particularly chloroform in potable water revealed that direct aqueous injection results were consistently higher than those obtained by gas sparging. A similar trend was shown for chlorodibromomethane. In general, the direct aqueous injection method indicated chloroform and bromodichloromethane concentrations 1.5 and 2.2 times higher respectively, than the values obtained by gas sparging. These results suggested to Nicolson et al. [139] that actual chlorinated natural water samples, as opposed to synthetic standards, contain nonvolatile haloform precursors which are injected in determinations by direct injection but not by gas sparging. If the additional quantities of haloforms observed by the direct aqueous injection method are produced during analysis from non-volatile halogenated organic compounds, then in a given sample the difference should be measurable by analysing prepurged samples by direct aqueous injection. Earlier studies by Nicolson and Meresz [132] had indicated that a gas chromatograph equipped with a Poropak Q column and an electroncapture detector could be used for the determination of haloforms in dilute aqueous solution by direct aqueous injection. It was shown that the determination of the haloforms (chloroform, bromodichloromethane and chlorodibromomethane) as well as carbon tetrachloride, trichloroethylene and tetrachloroethylene could be achieved near the 1µg L-1 level using 9µL injection volumes. Since no preconcentration was required, this technique was easily automated, allowing the analysis of up to 60 samples per day. However, before adopting this method for routine monitoring, it was essential to compare it with the published gas sparging procedure.
Haloforms
389
Analysis of standard aqueous haloform solutions showed excellent agreement between the two methods. However, when the two methods were compared on samples of chlorinated water obtained from treatment plants a different picture emerged. The direct aqueous injection method gave haloform values consistently higher than those obtained by gas sparging. The differences in data were consistent with the location of sampling sites. A study of this problem revealed that the two methods are measuring different parameters. Namely, the gas sparging technique quantitates only the free haloforms, while the direct aqueous injection method measures the total potential haloform concentration. The direct aqueous injection technique will produce a value which is the maximum haloform concentration which can be reached while the water is in the distribution system. When the free haloform content of the sample has been determined using both methods, the difference in the free and total potential haloform results gives an indication of the amounts of higher-molecular-weight haloform intermediates which cannot be detected by the gas sparging method. Quimby and Delaney [184] and Quimby et al. [185] determined trihalomethanes in potable water by gas chromatography with an atmospheric pressure microwave-emission detector. The organics are isolated by a purge and trap technique. This detector, in addition to being very sensitive, is also element selective, distinguishing between chlorine, bromine and iodine. The water sample is contained in a glass tube 6in (15cm) long. Helium or nitrogen purge gas, which has passed through a trap containing silver oxide, is heated at 200°C to oxidize any organic contaminants bubbled into the water sample. The organics purged from the water sample are desorbed from the Tenax GC trap into the gas chromatograph by attaching the end of the trap to a helium line and inserting the entire length of the trap into the injection port of the gas chromatograph. Carrier gas is then purged through the heated Tenax GC trap on to the gas chromatographic column. For 5mL water samples, detection limits of ca. 0.1µg L-1 are obtained for the four bromine- and chlorine-containing trihalomethanes, and the linear range extends to above the 100µg L-1 level. Similar results are also obtained for the other organohalides. Chlorine-, bromine- and iodineselective chromatograms for a standard aqueous solution containing several organohalides at the 25–30µg L-1 level are given in Fig. 5.11. Pfaender et al. [186] have compared purge and trap and direct injection gas chromatographic techniques for the determination of chloroform in potable water and found evidence for the presence of non-volatile precursors. The direct aqueous injection technique, employing a bypass valve to vent water and an electron-capture detector, gave consistently higher values for chloroform than the
390
Halogen-containing compounds
Fig. 5.11 Element-selective chromatograms from sparged standard solution, (a) Cl channel, (b) Br channel, (c) I channel. (Reprinted with permission from Quimby and Delaney [184]. Copyright (1979). American Chemical Society.)
purge method. Comparable results were obtained if the direct injection value after a 30min purge was subtracted from the beforepurged value. The nature of the residual measured by direct injection after purging was investigated and shown to be due to nonpurgeable intermediates that decompose within the injection port of the gas chromatograph to chloroform. The residual varied depending on the source of the water sample examined and the specific configuration of the chromatograph employed. The results indicated the need for caution in the interpretation of chloroform and other trihalomethane values, especially haloform potentials, generated by direct aqueous injection. Pfaender et al. [186] were unable to identify the chloroform precursor in potable water samples. They did conclude that the size of the residual measured by direct aqueous injection depends on the materials and configuration of the injection port of the chromatograph used. This raises doubts as to the usefulness of the residual haloforms measured by direct aqueous injection as an estimate of haloform formation potential. The relationship of the potential as determined by direct aqueous
Haloforms
391
injection and the haloforms formed during distribution of a finished drinking water is still largely unknown. Kirschen [102] has investigated the Environmental Protection Agency standard purge and trap method [187] for determining trihalomethanes and other halogenated volatiles in water. Kirschen [102] used a Varian Model 3700 gas chromatograph with Model 700A Hall electrolytic conductivity detector VISTA 401 chromatography data system. The 2m × 2mm glass column contained 1% SP 1000 on Carbopack-B (60–80 mesh). The instrument parameters were: • • • • • •
Reactor temperature 750°C. Hydrogen flow rate 30mL min-1. n-Propanol flow rate 0.6mL min-1. Detector base temperature 230°C. Range 100. GC conditions: carrier gas helium, 30mL min-1. Temperatures: column oven 45°C for 3min, 8°C min-1 to 220°C, hold for 2min; injector 150°C.
This method was used to check the concentrations of four halomethanes, chloroform, bromoform, dichlorobromomethane and dibromochloromethane in potable water samples. Relative percentage standard deviations were normally below 5%. A purging method has been described for the specific determination of low levels of methyl bromide fumigant in water [104]. The analysis was performed with a Hewlett Packard Purgatrator, mounted on a PTC (packed trap capillary) module, with two electron-capture detectors. The module was used in a Hewlett Packard Model 433 gas chromatograph. The essential instrumentation for methyl bromide determination consists of: a Purgatrator, a single-packed column and an electroncapture detector. This configuration is compatible with the Hewlett Packard 429, 430 and 433 gas chromatographs. Chiba and Haraguchi [188] determined trihalomethanes in microgram per litre amounts in potable water by gas chromatography– atmospheric pressure helium-microwave-induced plasma-emission spectrometry with a heated discharge tube for pyrolysis. The trihalomethanes are collected on a Tenax GC column by a gas purge and trap method. Purge closed-loop gas chromatography has been used [189] to determine trihalomethane in potable water. This method combines the technique of gas stripping and static headspace sampling. Recovery efficiencies of between 81 and 100% are obtained. Better quality chromatograms with faster handling were achieved using the purge closed-loop method than by the purge and trap technique.
392
Halogen-containing compounds
5.3.3.5 High-performance liquid chromatography This method has been used to determine non-volatile chlorinated hydrocarbons in potable water [190]. Reunanen and Kronfeld [191] describe a method for determination of volatile hydrocarbons in, among others, raw water and drinking water. The sample is extracted with petroleum ether and without further purification analysed by high-performance liquid chromatography with electron-capture detection. The levels of 10 volatile halocarbons in potable water were determined. Levels of trichloromethane in potable water were over 40 times higher than in the river water samples. 5.3.3.6 Miscellaneous Unexpected peaks appearing in the closed-loop stripping–gas chromatography gas chromatograms of potable water samples following the addition of bromoalkanes as internal standards have been shown to be due to chloralkanes of the same carbon number produced by conversion of bromoalkanes by free chlorine in the sample [192]. Miller and Uden [193] have studied the effects of chlorination parameters, such as reaction time, chlorine/carbon ratio, pH and source of humic material, on the formation of non-volatile organic chlorine compounds during the chlorination process. The process was followed both qualitatively and quantitatively. The quantitation procedure for haloforms in the chlorine–fulvic acid reaction consists of air-tight sampling vials, purge and trap techniques and gas chromatography with a microwave plasma-emission detector. Chlorine-selection fingerprint studies were undertaken to follow the formation of 17 major chlorinated products. Detailed data are presented on the formation of chloroform, dichloroacetic acid, trichloroacetic acid and chloral hydrate. Kissinger and Fritz [70] used the gas chromatographic technique to determine chloroform, bromodichloromethane and bromoform in chlorinated drinking water. They observed that the concentrations of these substances increased with the time of storage of the sample. Reichert and Lochtman [148] studied the appearance of haloforms in raw water and their formation during potable water treatment, as well as their elimination and the prevention of their formation. Techniques for the determination of organohalogen compounds are discussed, in particular, the extractive organic halogens and adsorbed organic halogen; methods are described and compared. A combination of these methods has been used to analyse surface waters in Germany. The adsorbed organic halogens are first determined by adsorption on activated carbon, followed by pyrolysis and microcoulometry. Parallel to these determinations the extractive organic halogens are determined according to the DIN regulation method, which
Haloforms
393
involves extraction with pentane–di-isopropyl ether, pyrolysis and determination of halogens in a condenser, by coulometry or nephelometry. 5.3.3.7 Preconcentration Active carbon has been used to preconcentrate chlorinated hydrocarbons from potable and waste waters prior to analysis [194, 195]. Ion-exchange resins have also been used to preconcentrate haloforms (Section 5.3.3.1). 5.3.4 Trade effluents 5.3.4.1 Solvent extraction–gas chromatography The pentane extraction capillary column gas chromatographic procedure described in Section 5.3.3.1 [96] has been applied to the determination of haloforms in chlorine-treated sulphate pulp mill effluents. Methylene dichloride (640µg L-1), chloroform (760µg L-1), CHClCCl (2.8µg L-1) and 3 CCl CCl (1.4µg L-1) were found in these effluents. 2
2
5.3.4.2 Headspace analysis The headspace method described by Dietz and Singley [82] and discussed in Section 5.3.1.3 has been applied to the determination of haloforms in trade effluents. 5.3.5 Sewage 5.3.5.1 Solvent extraction–gas chromatography De Leer [196] has described a procedure for the determination of trihaloforms (chloroform, bromodichloromethane, dibromochloromethane, bromoform) in secondary and tertiary treated sewages and in chlorinated sewage. The haloforms are extracted into pentane followed by gas chromatographic separation and quantification by electron-capture detection. The sensitivity of this procedure was more than adequate (less than 2µg L-1) for these types of sample. 5.3.6 Swimming pool waters 5.3.6.1 Solvent extraction–gas chromatography A gas chromatographic procedure for determination of volatile organochlorine compounds in swimming pool water has been described
394
Halogen-containing compounds
[197]. A column packed with Carbopack B with 0.8% SE-30 as the mobile phase is used, the sample being extracted by n-pentane in the sampling container. Retention times are given for a range of chlorinated and bromo-chloro derivatives. The method was applied to samples from enclosed public swimming baths, originally filled with water from the mains, and to medicinal baths, filled with thermal spring water containing detectable amounts of bromine in solution. While chloroform was the major chlorinated product in the enclosed baths, dibromochloromethane was the principal product in the baths containing thermal spring water, while chloroform was present only in very small quantities. 5.4 Haloaromatic compounds 5.4.1 Natural waters 5.4.1.1 Gas chromatography Kapila and Aue [198] have studied the determination of hexachlorobenzene using electron-capture gas chromatography. The technique involves routing of gas chromatographic peaks of chlorinated hydrocarbons to a built-in flow-through reactor where they are partially dechlorinated in carrier gas doped with hydrogen, over a nickel catalyst, and the reaction products flow on to a second column for separation. Oliver and Bothen [199] discuss a capillary gas chromatographic procedure which can be used, after sample preconcentration, to identify and quantify 12 chlorinated benzenes in water samples. Preconcentration factors of 1000 with a small column of macroreticular resin, and 2500 with liquid–liquid extraction using pentane, were achieved. The pentane extraction technique is preferred because of its simplicity. Well-coated open tubular glass capillary columns were used for gas chromatography with a nickel60 electron-capture detector. Using both preconcentration techniques more than 80% of all the chlorobenzenes were recovered. Detection limits in water for the pentane extraction technique vary from about 1ng L -1 for dichlorobenzene to about 0.01mg L -1 for hexachlorobenzene. Italia and Uden [200] used electron-capture gas chromatography to compare the volatile halogenated compounds formed in the chlorination of humic acid. Ryan et al. [201] used a helium discharge detector for the quantisation of volatile haloaromatics in natural waters. Down to 20pg of pdibromobenzene could be determined.
Polychlorobiphenyls
395
5.4.1.2 Liquid chromatography Di Corcia and Samperi [202] determined down to 0.1µg L -1 of chloroaniline in environmental waters. Acetonitrile was used as extractant with two extractors in tandem. 5.4.1.3 Miscellaneous Stetter and Cao [203] used a gas sensor and a permeation apparatus for the determination of 1–5mg L-1 of chlorinated hydrocarbons in water. 5.5 Chlorinated alkyl naphthalenes 5.5.1 Trade effluents 5.5.1.1 Miscellaneous Bjorseth et al. [204] have determined these substances in bleach works effluents by a combination of glass capillary gas chromatography, gas– liquid chromatography–mass spectrometry and neutron activation analysis. 5.6 Chlorinated polyaromatic compounds 5.6.1 Waste waters 5.6.1.1 High-performance liquid chromatography–gas chromatography Oyler et al. [205] have determined aqueous chlorination reaction products of polyaromatic hydrocarbons in amounts down to micrograms per litre using reversed-phase high-performance liquid chromatography– gas chromatography. The method involves filtration through a glass microfibre filter and concentration on a high-performance liquid chromatography column. The polyaromatic hydrocarbon material is then eluted using an acetonitrile–water gradient elution technique. The fractions are injected separately on to a gas chromatographic column equipped with a photionization detector. 5.7 Polychlorobiphenyls 5.7.1 Natural waters Gas chromatography, possibly coupled with mass spectrometry, predominates as a method of choice for polychlorobiphenyls (PCBs).
396
Halogen-containing compounds
They have been prepared industrially since 1929 by chlorination of biphenyl with anhydrous chlorine using either iron filings or iron (III) chloride as a catalyst. The product obtained is a complicated mixture of several PCBs. In the UK, PCBs are marketed as Arochlors by Monsanto. All Arochlors are characterized by a four-digit number; the first two digits represent the type of molecule (e.g. 12 represents biphenyl, 54 terphenyl and 25 and 44 are mixtures of biphenyl and terphenyl); the last two digits give the percentage by mass of chlorine, e.g. Arochlor 1260 is a 12-carbon system with 60% m/m of chlorine. PCBs are sold under a variety of trade names, of which Arochlor is one. The following is a list of the principal trade names used for PCB-based dielectric fluids which are usually classified as Askarels: Arochlor (UK, USA), Pyoclor (UK); Inerteen (USA); Pyanol (France); Clophen (Germany), Apirolio (Italy); Kaneclor (Japan); Solvol (Russia). Other names were used for PCB products intended for different applications but are no longer in current use; these include: Santotherm FR (UK, prior to 1972 for heat transfer); Therminol FR (USA, prior to 1972 for heat transfer); Pydraul (USA, prior to 1972 for hydraulic applications); Phenoclor (France) and Fenclor (Italy). The trade names Santotherm, Therminol and Pydraul are still in use but they are now used to refer to non-chlorinated products. 5.7.1.1 Gas Chromatography Webb and McCall [206] isolated 27 isomers found in Arochlors 1221, 1242 and 1254 by preparative gas chromatography and identified them by comparison of their retention data and infrared spectra with those of known synthetic compounds. Analytical gas chromatography was carried out on a support coated open-tubular column (100ft × 0.2in (30.5m × 5mm)) of SE-30 at 190°C with helium as carrier gas (6.5mL min-1) and flame ionization detection. Bauer [207] has described a gas chromatographic procedure for the determination of PCBs in water. The PCBs were extracted from water with hexane, and the extract was dried and concentrated before gas chromatographic analysis. Extraction and concentration of PCBs were also effected by the use of slow filtration through sand or algae columns. The sand was extracted with 60% acetone–hexane, acetone was removed by washing the extract with water, and the washed hexane was passed through a column filled with layers of sodium sulphate, alumina and Florasil. The eluate was analysed by gas chromatography. Algal material was dried, ground with sand, and treated in a manner similar to that used for sand alone. The gas chromatography was carried out on glass columns (3m × 2.8mm) packed with 2.5% QF-1, 2.5% silicone rubber and 0.5% Epikote 1001 on
Polychlorobiphenyls
397
Chromosorb W AW-DCMS (80–100 mesh) and operated at 200°C with nitrogen (90mL min-1) as carrier gas and an electron-capture detector. Berg et al. [208] separated PCBs from chlorinated insecticides on an activated carbon column prior to derivatization and gas chromatographic separation on a capillary column. Separation is based on the observation that PCBs adsorbed on activated charcoal cannot be removed quantitatively with hot chloroform but can be with cold benzene. Insecticides of the DDT group and a variety of others (e.g. ?BHC, Aldrin, Dieldrin, Endrin, and Heptachlor and its epoxide) can be eluted from the charcoal with acetone–ethylether (1:3). Typical recoveries from a mixture of p,p’-DDE (1,1-dichloro-2,2-bis-(4chlorophenyl)ethylene), p,p’-TDE, and o,p’-DDT and a PCB (Arochlor 1254) by successive elution with 90mL of 1:3 acetone–ether and 60mL of benzene were 91, 92, 92, 94 and 90%, respectively. Identification and determination of the PCBs was effected by catalytic dechlorination to bicyclohexyl or perchlorination to decachlorobiphenyl, followed by gas chromatography. For bicyclohexyl, gas chromatography was carried out on a column (8ft × 0.25in (2.4m × 6mm)) of 10% DC-710 on Chromosorb W, operated at 90°C for 2.5min then temperature programmed at 10°C per min, with flame ionization detection. For decachlorobiphenyl, the column (2ft × 0.125in (0.6m × 3mm)) is 5% SE-30 on Chromosorb W, operated at 215°C, with nitrogen as carrier gas and a tritium detector. Beezhold and Stout [209] studied the effect of using mixed standards on the determination of PCBs. Mixtures of Arochlors 1254 and 1260 were used as comparison standards and gas chromatograms of these mixtures were compared with those obtained from a hexane extract of the sample after clean-up on a Florasil column. Polychlorinated biphenyls were separated from DDT and its analogues on a silica gel column activated for 17h and with 2% (w/w) of water added. The extracts were analysed on a silanized glass column packed with 5% DC200 and 7.5% QF-1 on Gas Chrom Q (80–100 mesh) operated at 195°C with nitrogen as carrier gas (50–60mL min-1) and a tritium detector. Glass-wall coated open tubular capillary columns have been used for the gas chromatography of PCBs [210, 211]. Figure 5.12 shows a chromatogram obtained by this technique [210] for a mixture of Arochlors in hexane. The column comprised 25mL × 0.25mm of WCOT glass CPtm Sil 7. It can be seen that 0.1ppm of these substances is easily detected. Schulte and Acker [211] gas chromatographed PCBs with a glass capillary column at temperatures up to 320°C. They used a 60m column impregnated with SE-30-SC and helium as carrier gas with a flame ionization detector. The column was maintained briefly at 80°C after injection of the sample, heated to 180°C and then temperature programmed to 260°C at 20°C per minute.
398
Halogen-containing compounds
Fig. 5.12 Chromatogram of Arochlor 1260. (Copyright (1979) [210] Chromopak International B.V.)
Brinkmann et al. [212, 213] used a silica gel column which elutes the higher chlorinated PCBs in the normal phase. This system produced a reasonable separation of the lower chlorinated PCBs present predominantly in the commercial mixture Arochlor 1221, but was less efficient in separating the more highly chlorinated PCBs present in Arochlors 1254 and 1260. Dunn et al. [214] propose a technique for the quantitative determination of constituents in complex mixtures characterized by gas chromatography data using partial least squares in latent variables. The technique was applied to the gas chromatograms of Arochlor 1242, 2148, 1252 and 1260. Two SIMCA pattern recognition analyses were carried out: classification of an unknown sample of a specific Arochlor, and a calibration in which the relative amounts of specific Arochlors were estimated in a classified sample. Other references to the gas chromatography of PCBs in natural water samples are reviewed in Table 5.13.
Polychlorobiphenyls
399
Table 5.13 Chromatography of PCBs in natural water
5.7.1.2 Gas chromatography–mass spectrometry The earliest reference to the application of this technique occurrs in 1971 when Stalling and Huckins [225] discussed gas chromatography– mass spectrometry characterization of Arochlors and 36Cl labelling of Arochlors 1248 and 1254. These workers mention the separation of PCBs from polyterphenyls. Ahnoff and Josefsson [226] carried out confirmation studies on PCBs from river waters using mass fragmentography, a form of gas chromatography–mass spectrometry in which the mass spectrometer is focused at one or more individual mass numbers while the components emerge from the column. This technique is a more specific and sensitive technique than electron-capture gas chromatography for detecting and identifying traces of PCBs in river waters. The specificity is further increased by using the known intensity ratios of the isotopic peaks of chlorine-containing species. Webb and McCall [227] determined the weight of PCBs producing each peak in chromatograms of several Arochlors by gas chromatography–mass spectrometry and the use of an electrolytic conductivity detector. Electron-capture detector response factors were derived, and these were applied in association with rules for the division of chromatograms to the quantitative analyses of environmental samples containing one or more Arochlors. Eichelberger et al. [228] applied gas chromatography–mass spectrometry with computer-controlled repetitive data acquisition from selected specific ions to the analyses of PCBs in lake sediments. The PCB mixtures were separated by gas chromatography at 180°C in a coiled glass column (6ft
400
Halogen-containing compounds
× 0.78in (1.8m × 20mm)) packed with 1.5% OV-17 plus 1.95% QF-1 on GasChrom Q (100–120 mesh), with helium (30mL min-1) as carrier gas. Effluent is passed via a glass jet enrichment device into a quadrupole mass spectrometer, controlled by a minicomputer in such a way that only selected ions of specific m/e pass through the quadrupole field. There is a substantial gain in sensitivity, without loss of qualitative information contained in the complete mass system. This technique provides a basis for a sensitive qualitative and quantitative (from ionabundance chromatograms obtained from subset scanning) analysis for PCBs. Karlruber et al. [229] and Ahnoff and Josefsson [230] have discussed the application of mass fragmentography to the detection of traces of PCBs and herbicides in water. Voyksner et al. [231] compared gas chromatography–high-resolution mass spectrometry and mass spectrometry–mass spectrometry for the detection of PCBs and tetrachlorodibenzofuran in water. Pellazari et al. [232] have reviewed advances in the detection, identification and quantification of PCBs. Areas covered included highresolution gas chromatography, using capillary columns, and detection systems (electron capture, negative-ion chemical ionization mass spectrometry, selected ion monitoring and pulsed positive-ion/negativeion chemical ionization mass spectrometry). The availability of chlorobiphenyl isomer standards is discussed. Tanabe et al. [233] have described a method for the determination of specific tetra-, penta- and hexachlorinated PCB isomers, related to the highly toxic 2,3,7,8-tetrachlorodibenzodioxin. Stages consist of alkaline extraction, clean-up and fractionation on activated carbon followed by high-resolution gas chromatography with electron-capture detection and mass spectrometric identification of individual isomers. 5.7.1.3 Thin-layer chromatography de Vos and Peet [234] applied mixtures of PCBs in hexane solution (20µL) to a 0.25 mm layer of Kieselgel G previously saturated with light petroleum containing 8% of liquid paraffin and dried. The chromatograms were developed ( × 3) with acetonitrile–acetone– methanol–water (20:9:20:1) previously saturated with liquid paraffin. After drying in air, the plates were sprayed with 0.85% ethanolic silver nitrate and exposed to UV radiation for detecting the spots (black on a white background). Sackmauer et al. [235] determined PCBs in water by thin-layer chromatography on silica gel plates impregnated with 8% paraffin oil. As a mobile phase a mixture of acetonitrile, acetone, methanol and water (20:9:20:1) was used. For detection a solution of silver nitrate
Polychlorobiphenyls
401
and 2-phenoxyethanol was used, followed by irradiation with UV light. The detection sensitivity for Arochlor 1243 is 0.5-1 .0µg. 5.7.1.4 High-performance liquid chromatography Albro and Fishbein [219] determined the retention indices on six stationary phases for some mono-, di-, tri-, tetra- and hexachlorobiphenyls. They confirmed the additivity of the half-retention values in predicting retention indices. Brinkman et al. [212, 213] used a silica gel column which elutes the higher chlorinated PCBs in the normal phase. This system produced a reasonable separation of the lower chlorinated PCBs present predominantly in the commercial mixture Arochlor 1221 but was less efficient in separating the more highly chlorinated PCBs present in Arochlors 1254 and 2160. Kaminsky and Fasko [236] investigated the potential of reversed phase liquid chromatography to the analysis of PCB mixture in environmental samples. They used mixtures of water and acetonitrile as the mobile phase to achieve analysis of 49 different PCBs and of samples of Arochlor 1221, 1016, 1254 and 1260. Some of the major constituents of the Arochlor are tentatively assigned. For Arochlor 1221 (21% by weight of chlorine), 31 components were distinguishable chromatographically. The seven major components were tentatively identified and quantitated. 3-chlorobiphenyl is not completely resolved from an excess of 2-chlorobiphenyl and a trace amount of the former may be present and thus increase the reported value for the latter. It is apparent that the patterns of minor components of Arochlor 1221 closely resemble those of the major components of Arochlor 1016 and 1254. Various other workers have discussed the high-performance liquid chromatography of PCBs [212, 213, 236–239]. Petrick et al. [239] used high-performance liquid chromatography to remove aliphatic compounds and polyaromatic hydrocarbons from an n-hexane extract of a natural water sample prior to its analysis by gas chromatography for PCB congeners. 5.7.1.5 Polarography PCBs, chlorinated insecticides and polychlorinated naphthalenes and benzenes have been identified voltammetrically [240] using threeelectrode potentiostatic control circuitry with interruptable linear voltage sweep control and a normal voltage scan rate of 142mV s-1. The reference electrode was a standard calomel electrode, the auxiliary electrode was a platinum wire and the stationary electrode was a mercury-coated platinum electrode. Dimethyl sulphoxide was used as solvent with 0.1M
402
Halogen-containing compounds
tetraethylammonium bromide as the supporting electrolyte, and nitrogen was used for deaeration. Neely [241] determined PCBs in water in Lake Michigan. Kopanica [242] applied polarographic and voltammetric techniques for the analysis of trichlorobiphenyl in natural waters using fast scan differential pulse polarography with a hanging mercury drop electrode. 5.7.1.6 Enzyme immunoassay Aga et al. [243] determined Arochlor and its sulphonic metabolite in natural waters using solid-phase extraction and enzyme-linked immunosorbent assay. 5.7.1.7 Miscellaneous Polyurethane foam has been used as a liquid–liquid partitioning filter for the concentration of PCBs from water samples [244, 245]. Ahnoff and Josefsson [246] tested various clean-up procedures for PCB analysis on river water extracts. Extracted water samples were divided into a number of portions and used in testing the following clean-up treatments: with sulphuric acid, with sulphuric acid and activated Raney nickel, with a Florasil column, with a Florasil column and activated Raney nickel, and with a Florasil column and potassium hydroxide. The results of subsequent gas chromatographic analyses indicate the Florasil column procedure is more effective in removing contaminants than the sulphuric acid procedure. However, neither procedure could remove sulphur, which could be removed by the activated Raney nickel or potassium hydroxide procedures. The US Environmental Protection Agency [247] and the Inland Waters Directorate, Canada [248] have described a method for the determination of PCBs in water. 5.7.1.8 Preconcentration PCBs are often extracted from water by solvent extraction. However, the relatively small sample size, ca. 1-2L, that can be handled restricts the quantitation limits of PCB analyses. Ion-exchange resins Various methods have been described for the extraction and recovery of PCBs prior to gas chromatographic analysis. Coburn et al. [249] have shown that XAD-2 macroreticular resin can be successfully
Polychlorobiphenyls
403
used to analyse PCBs in 2L natural water samples fortified at the 250ng L1 level. Amberlite XAD-2 and XAD-4 macroreticular resins have been used by several workers (Chriswell et al. [250], Musty and Nickless [251, 252]) to preconcentrate PCBs. Polyurethane plugs Porous polyurethane foam has also been used to preconcentrate water samples. Gesser et al. [253] found that the compounds could be adsorbed on a column composed of two polyurethane plugs (each 38mm × 22mm) inside a glass tube. The sample water was poured through at 250mL min -1, then the plugs were removed and squeezed free from water; the PCBs were then extracted by treating the plugs with acetone and hexane. The concentrated extract was analysed by gas chromatography on a glass column (6ft × 0.25in (1.2m × 6mm)) packed with Chromosorb W HMDS supporting 2% SE-30 and 3% QF-1 and operated at 200°C; the carrier gas was helium and a 63 Ni electron-capture detector was used. In tests on 20µg of PCBs added to 1L of water, the recovery (based on measurements of 13 peaks on the chromatogram) ranged from 18.2 to 19.6µg. Bedford [254] also used polyurethane-foam plugs to extract PCBs from natural waters; these results indicated that Arochlor is probably adsorbed on to small particles in unfiltered lake water which can pass through the foam. Although this makes the foam method somewhat impractical for enriched and turbid waters, it is still a valuable technique for clear water containing low quantities of PCBs. Other materials that have been used to preconcentrate PCBs include: charcoal [250], Carbowax–undecane on Chromosorb W [252], macroreticular resins [250, 252, 255], cellulose triacetate [256] and covalently bonded silica gel (with octadecyl groups) [257]. 5.7.2 Sea water 5.7.2.1 Gas chromatography Elder [258] determined PCBs in Mediterranean coastal waters by adsorption on the XAD-2 resin followed by electron-capture gas chromatography. The overall average PCB concentration was 13ng L-1. 5.7.2.2 High-performance liquid chromatography Petrick et al. [239] used high-performance liquid chromatography to remove aliphatic compounds from estuary waters prior to determining PCBs by gas chromatography.
404
Halogen-containing compounds
5.7.2.3 Miscellaneous Sullivan et al. [259] studied the loss of PCBs from sea water samples during storage. 5.7.3 Potable waters 5.7.3.1 Gas chromatography Albro and Fishbein [219] in 1972 carried out early work on the quantitative and qualitative analaysis of PCBs using gas chromatography with flame ionization detection. Webb and McCall [227] studied the question of quantitative PCB standards for electron-capture gas chromatography. Le’Bel and Williams [260, 261] found that low procedural blank values, equivalent to 0.04ng L-1 for a 200L potable water sample were attainable only by using doubly distilled solvents and by exhaustive washing of all reagents and glassware with these solvents. Gas chromatographic analysis of concentrated extracts of potable water samples without Florasil column clean-up gave off-scale peaks at instrument settings suitable for low nanogram per litre PCB analysis. However, fractionation of the extract by Florasil column chromatography gave a PCB fraction sufficiently clear of interfering organics to permit PCB analysis at 1–10ng L-1. The gas chromatographic analyses were done on a relatively short (11m) capillary column, thus providing a compromise of short analysis time and good resolution with minimum interferences from other substances present in the water extract. Recoveries of Arochlors 1232, 1016, 1242 and 1254 from water samples fortified at the equivalent of 1 and 10ng L-1 for a 200L sample ranged from 86 to 99% at the 10µg L-1 level and from 91 to 110% at the 1µg L-1 level. The detection limit was estimated to be ca. 0.04ng L-1 of Arochlor 1016 from this source of water with Arochlors 1232, 1242 and 2154 having similar levels of detection. When the method was applied to potable water from a river source, the interference in the gas chromatogram from other organic compounds present in the sample made quantitation difficult at the 1ng L-1 level. Arochlor 1254 would still be detectable at the 1ng L-1 level but nanogram per litre levels of Arochlors 1232, 1016 and 1242 would not have been distinguishable from the background peaks. However, at the 10ng L-1 level, it was possible to distinguish peaks due to Arochlor 1232, Arochlor 1242 and Arochlor 1016 from the background chromatogram due to other organics. It is concluded that the XAD-2 macroreticular resin method can be used to analyse large volumes of potable water for low
Polychlorobiphenyls
405
nanogram per litre levels of PCBs. The detection limits, however, are limited by the interference from other organics in the water sample but are typically in the 1–10ng L-1 range. The analysis of 200L potable well water samples, suspected of contamination of Arochlor 1016, gave results in the range less than 0.05–0.02ng L -1 PCB (calculated as Arochlor 1016). The interfering compounds were minimal except for an amplified series of peaks similar to the two-fold concentration procedural blank. The higher background was due to analysis before the method was fully optimized for minimum interference. Quantitation of the Arochlor 1016 was done by comparing peaks from the interference-free region between retention times 3–7min, where several of the Arochlor 1016 components were eluting. When the method was applied to potable water from a river source, the interference in the gas chromatogram from other organic compounds present in the sample made quantitation difficult at the 1ng L-1 level. Arochlor 1254 would still be detectable at the 1ng L-1 level but 1ng L-1 levels of Arochlors 1232, 1016 and 1242 would not have been distinguishable from the background peaks. However, at the 10ng L-1 level, it was possible to distinguish peaks due to Arochlor 1232, Arochlor 1242 and Arochlor 1016 from the background chromatogram due to other organics. 5.7.4 Trade effluents. 5.7.4.1 Gas chromatography Various workers have described methods (gas chromatographic) for determining PCBs in paper mill effluents. The method described by Delfino and Easty [262] is capable of detecting down to 2µg L-1 PCB. Easty and Wabers [263, 264] have studied the effect of suspended solids on the determination of Arochlor 1242 in paper mill effluents. 5.7.4.2 Column chromatography Lopez-Avila et al. [265] evaluated Environmental Protection Agency column chromatographic method no. 8080 for the determination of chlorinated insecticides and PCBs in hazardous waters. They found that the Florasil column procedure recommended by the EPA did not separate organochlorine insecticides from PCBs and developed an alternative procedure using a silica gel column and a sulphur clean-up for achieving this separation. The revised procedure had detection limits of 0.02–0.09 and 0.5–0.9µg L-1, respectively, for organochlorine insecticides and PCBs.
406
Halogen-containing compounds
Easty and Wabers [263, 264] have studied the effect of suspended solids on the determination of Arochlor 1242 in paper mill effluents. 5.7.5 Sewage effluents 5.7.5.1 Gas chromatography Workers at the US Environmental Protection Agency [266] have evaluated protocols for chlorinated insecticides and PCBs in raw waste water and sewage effluents. They concluded that the gas chromatographic method performed satisfactorily at the parts per billion level. 5.7.5.2 Gas chromatography–mass spectrometry Skinner et al. [267] showed for the first time the application of mass fragmentography to residue analysis, namely the determination of PCBs along with DDE in extracts of a sewage effluent. 5.7.5.3 Miscellaneous Kaiser and Wong [332] studied the bacterial degradation of Arochlor 1242 in water and identified aliphatic and aromatic hydrocarbon metabolites using gas chromatography and mass spectrometry. No evidence was found for chlorine-containing metabolites. 14C-labelled PCBs have been used [268, 269, 272] to ascertain the fate of PCBs in activated sludge in a sewage works. It was found that tri- and pentachlorobiphenyl were practically unaffected by bacterial attack and were partitioned, in unchanged form, between the water and the sludge, a major portion being retained by the sludge. Furukawa et al. [269] studied the effect of chlorine substitution on the biodegradability of 31 different PCBs. Their biodegradability using Alcaligenes and Acinetobacter, showed that degradation varied inversely with chlorine substitution. PCBs with chlorine in the ortho position showed poor degradability; those with all chlorine atoms on a single ring degraded faster than those with the same number on both rings; preferential ring fission occurred with non-chlorinated or less chlorinated rings; and only in the degradation of 2,4,6trichlorobiphenyl were significant differences observed between the two organisms. Gaffney [270] showed that PCBs were present in certain municipal sewages. He showed that various chlorobiphenyl isomers are produced during chlorination of sewage and sewage effluents.
Chlorophenols
407
5.8 Chlorophenols 5.8.1 Natural waters 5.8.1.1 Spectrophotometric method Carr et al. [271] give details of a simple procedure for the determination of pentachlorophenol in water. Although this method is not as sensitive or specific as gas chromatography, the number of potential interfering compounds is limited and it is useful for routine monitoring. 5.8.1.2 Gas chromatography Earlier references to the application of gas chromatography to the determination of Chlorophenols in natural waters are summarized in Table 5.14. Rudling [273] determined down to 100µg L-1 pentachlorophenol (PCP) in water by an electron-capture gas chromatographic method. Hexane (0.5mL) and fresh acetylation reagent (pyridine (2mL) plus acetic anhydride (0.8mL) stored in the cold) (40µL) are added to the combined aqueous extracts and shaken for 1min. The hexane phase is analysed by gas chromatography on a glass column (1m × 1.5mm) packed with 5% of QF-1 on Varaport 30 (100–120 mesh) operated at 150°C with nitrogen as carrier gas (25mL min-1).
Table 5.14 Determination of Chlorophenols (PCP) in natural water by gas chromatography
408
Halogen-containing compounds
Renberg [274] has used an ion-exchange technique for the determination of chlorophenols and phenoxyacetic acid herbicides in water. The water samples are mixed with Sephadex QAE A-25 anion exchanger and the adsorbed materials are then eluted with a suitable solvent. The chlorinated phenols are converted into their methyl ethers and the chlorinated phenoxy acids into their methyl or 2-chloroethyl esters for gas chromatography. Renberg [274] recommends the use of ?-BHC (lindane), DDE or DDT as a gas chromatographic internal standard. The relative retention times of the derivatives corresponding to these internal standards are shown in Table 5.15. The detection limits for the different substances for 1L of a water sample are 0.0001–0.1ppb. Chriswell and Cheng [275] have showed that chlorophenols and alkylphenols in the ppb to ppm range in natural waters and treated drinking water can be determined by sorption on macroporous anionexchange resin, elution with acetone and measurement by gas chromatography. These workers describe techniques preventing phenol losses caused by chlorination, oxidation and other reactions during their determination. Common inorganic ions and many organic substances cause no interference; neutral organics that are retained by the resin can be removed by a methanol wash. Phenols are effectively Table 5.15 Retention times relative to ?-BHC, p,p’-DDE and p,p’-DDT
Source: Reprinted with permission from Renberg [274]. Copyright (1974) American Chemical Society.
Chlorophenols
409
retained from water when samples of water containing low concentration of phenols are made basic and passed through a column containing an anion-exchange resin. The basis of the method is as follows. Phenols are taken up as phenolate ions by passing an alkaline water sample through a column of A-26 anion-exchange resin in the hydroxyl form. Any neutral organic compounds retained by the resin are removed by washing with alkaline methanol. Phenolate ions continue to be held by the resin during this washing step and are then converted to the molecular form by washing the column with aqueous hydrochloric acid. The phenols are subsequently eluted from the column with acetone–water. The hydrochloric acid and acetone–water effluents are each extracted with methylene chloride. The organic phases are concentrated by evaporation and the phenols are separated by dual-gas chromatography. Hoben et al. [277] has described a gas chromatographic technique for determining 0.1ppb pentachlorophenol in water. Configuration of the identity of the chlorophenol was provided by gas chromatography– mass spectrometry. In this method the pentachlorophenol is extracted from the acidified sample with n-hexane and then re-extracted into a borax solution. It is then acetylated by extracting with n-hexane containing acetic acid anhydride and pyridine. The resulting pentachlorophenyl acetate is analysed by gas chromatography using an electron-capture detector. The extract procedure gave 91–98% recovery of pentachlorophenol from water samples. Korhonen and Knuutinen [279] studied the gas chromatography of chlorinated phenols and catechols using a non-polar SE-30 quartz capillary column. A wide range of degrees of chlorination were analysed with flame ionization detection. The elution order of the compounds was confirmed and the relative retention times are presented. While all the compounds could be separated without derivatization, simultaneous determinations resulted in overlapping peaks. Retention times tended to be shorter than those for corresponding acetate esters. A stronger ortho-effect was observed in the catechols. Optimal conditions for the detection of trace quantities of chloro, nitro and methyl phenols in 0.4–1.0mL water samples have been described [280]. Phenols were extracted with toluene, derivatized with heptafluorobutyricanhydride and separated and quantified by electron-capture gas chromatography or flame ionization gas chromatography. Detection limits for phenol, chlorophenols, p-cresol, dimethyl and nitrophenols were 0.01–0.20µg L-1 and 1.6µg L-1 for 2,4dinitrophenol.
410
Halogen-containing compounds
5.8.1.3 Thin-layer chromatography Various workers [281–284] have used this technique for determining pentachlorophenol and other chlorophenols in water samples. Thielemann and Luther [284] separated the chlorophenols on Kieselgel G plates with benzene as solvent. The spots were located by spraying with diazotized sulphanilic acid solution or with a mixture (1:1) of 15% ferric chloride solution and 1% K Fe(CN) solution. 3
6
5.8.1.4 High-performance liquid chromatography Column chromatographic techniques have been described for the determination of pentachlorophenol and other chlorophenols, but these methods are not suitable for the determination of low levels. The best method for determining pentachlorophenol is conversion into the methyl ether followed by analysis using gas chromatography with an electroncapture detector, or gas chromatography coupled with mass spectrometry [277]. Both of these methods require an extensive amount of pretreatment and highly trained personnel for the operation of the equipment. Ervin and McGinnis [285] attempted to overcome this problem by developing a high-performance liquid chomatographic method for determining in water low concentrations of pentachlorophenol and chlorinated impurities that occur in the technical grade material such as 2,3,4,6-tetrachlorophenol, mono-, di- and trichlorophenols, octa-, heptaand hexachlorodibenzo-p-dioxins, and a variety of other polychlorinated aromatic compounds. The method involves chloroform extraction of acidified waste water samples and rotary evaporation without heat. After redissolving in chloroform the samples were analysed directly by high-performance liquid chromatography on a microparticulate silica gel column. A number of solvent combinations are possible and 98:2 cyclohexane–acetic acid is preferred. The minimum detectable concentration is 1ppm (without sample concentration) and the coefficient of variation is 1– 2%. The type of separation achieved with a microparticulate silica gel column is shown in Fig. 5.13. The first peak, as determined by gas chromatographic–mass spectrometric analysis, consisted of a complex mixture of polychlorinated compounds, including octa-, hepta- and hexachlorodibenzo-p-dioxins as well as a mixture of products including 2,4,6-trichlorophenol. The third peak was mainly 2,3,4,6tetrachlorophenol and the fourth peak was pentachlorophenol.
Chlorophenols
411
Fig. 5.13 Separation of technical pentachlorophenol using cyclohexane–acetic acid (98:2) as the eluting solvent. Peaks: 1, mixture of dioxins and other polychlorinated products; 2, mixture of chlorinated phenols including trichlorophenol; 3, tetrachlorophenol; 4, pentachlorophenol. (Reprinted with permission from Ervin and McGinnis [285]. Copyright (1980) Elsevier Science Publishers.)
5.8.1.5 Ion chromatography Brandt and Kettrup [286] determined organic group parameters AOCL, AOBr and AOS in water by ion chromatography detection. Model compounds studied included 4-chlorophenol, 4-bromophenol and thiobenzamide. 5.8.1.6 Polarography Pentachlorophenol has been determined in water in amounts down to 0.3mg L-1 by differential pulse polarography [287]. 5.8.1.7 Miscellaneous Boyle et al. [288] have studied the degradation of pentachlorophenol in a simulated lentic environment. Ingram et al. [289] used a mass spectrometric isotope dilution technique to determine approximately 0.2µg pentachlorophenol in water with a relative standard deviation of 8%. Malissa et al. [290] described methods for the extraction and enrichment of chlorophenols from natural water for their
412
Halogen-containing compounds
determination by capillary gas chromatography–Fourier transform infrared spectroscopy. Derivatization was used to improve the chromatographic detection of phenols. This process reduced the differentiating power inherent in spectrometric gas chromatography detectors by introducing functional groups common to all environmental samples. The separation and identification of phenol and 19 chlorinated phenols from spiked river water without derivatization is reported. The method used capillary gas chromatography and Fourier-transform infrared spectroscopy, with an SE54 fused silica WCOT column. An on-column injection technique with a thermodesorption and cold trap injector with solvent removal was used. Mean recovery of chlorophenols from river water samples at concentrations of 10µg L-1 was in the 90% range. 5.8.1.8 Preconcentration Lipidex 500 [291] (1,4-dimethacryloxy methyl) naphthalenedivinyl benzene copolymer [292], activated carbon [293], XAD-4 resin [293] and C silica [294] have all been studied as solid adsorbents for the 18 preconcentration of chlorophenols prior to analysis by gas chromatographic [291, 294] and quartz spring balance techniques [293]. 5.8.2 Potable waters 5.8.2.1 Gas chromatography Chriswell and Cheng [295] have showed that chlorophenols and alkylphenols in the milligram to microgram per litre range in potable water can be determined by sorption on macroporous A-26 anionexchange resin, elution with acetone and measurement by gas chromatography (see also Section 5.8.1.2). These workers describe techniques preventing phenol losses caused by chlorination, oxidation and other reactions during their determination. Common inorganic ions and many organic substances cause no interference; neutral organics that are retained by the resin can be removed by a methanol wash. Phenols are effectively retained from water when samples of water containing low concentrations of phenols are made basic and passed through a column containing an anion-exchange resin. A number of neutral organic compounds (aliphatic alcohols, 2-phenoxyethanol, methyl cellulose, naphthalene, C –C aliphatic acids) are retained by A-26 resin. However, 4 8 none of these compounds affected recovery of phenols. They are eluted from a resin by basic methanol prior to elution of phenols and thus do not
Chlorophenols
413
interfere with the gas chromatographic determination of phenols. Carboxylic acids also cause no interference with the recovery of phenols using the procedure but can interfere with the gas chromatographic determinations of phenols on OV-17 columns by overlapping the phenol and cresol peaks. When a Tenax-GC column is used, acids elute well before phenols. Table 5.16 gives recoveries obtained by this procedure for potable waters spiked with various phenols (Fig. 5.14). Chriswell and Cheng [295] observed that low recoveries of phenols in chlorinated drinking water coincided with enhanced recoveries of chlorinated phenols. Addition of low concentrations of chloramine T to very dilute aqueous solutions of phenols caused similar results. Letting 3,5-dimethylphenol solutions prepared in chlorinated tap water stand for a few minutes resulted in 40% loss. New gas chromatographic peaks from extracts from this solution were positively identified by mass spectrometry as being the di- and trichlorodimethylphenols. Chlorination reactions during the determination of phenols can be prevented by addition of hydroxylamine hydrochloride. On real samples, however, it must be recognized that chlorination reactions may have occurred before the sample was taken. The recovery of pentachlorophenol was affected by the amount of reductants used to prevent oxidation and chlorination. The more reductant added, the lower its recovery. If the amounts of reductants specified in the recommended procedure are used,
Table 5.16 Recovery of phenols added to tap water
Source: Reprinted with permission from Chriswell and Cheng [295]. Copyright (1975) American Chemical Society. Notes a Using calibration curve: average of nine analyses except for 2-naphthol. b Using a single standard, a single analysis is reported to the closest 5%. The results in column five of the table show that the method is applicable for analysis of phenols at concentration levels as low as 5– 85ppb.
414
Halogen-containing compounds
Fig. 5.14 Separation of a standard mixture of phenols on a Tenax GC column. Peak order: (A) solvent impurity; (B) phenol; (C) o-cresol; (D) 3,5-dimethylphenol; (E) 4-chloro-3-methylphenol; (F) 2,4,6-trichlorophenol; (G) 2-napthol; and (H) pentachlorophenol. Separations were obtained on a 18 in × 1/4 in (0.46m × 6mm) in SS Tenax GC column. Temperature held at 190°C for 1min, then programmed at 10°C min-1 to 270°C and held at 270°C for 4min. (Reprinted with permission from Chriswell and Cheng [295]. Copyright (1975) American Chemical Society.)
pentachlorophenol losses will be negligible as will losses due to chlorination and oxidation of other phenols. Ashiya et al. [278] combine the phenols in the water with bromine, and measure the concentration of phenols and chlorophenols by electron-capture detection gas chromatography, obtaining their concentrations separately to the extent of 1µg L-1. Morgade et al. [296] have described an electron-capture gas chromatographic method for the determination of polyhalogenated phenols in chlorinated and unchlorinated potable water. The particular phenols studied were: 2,4-dichlorophenol (2,4-DCP); 2,3,5trichlorophenol (2,3,5-TCP); 2,4,5-trichlorophenol (2,4,5-TCP); 2,4,6trichlorophenol (2,4,6-TCP); 2,5-dichloro-4-bromophenol (2,5-DC-4-BP); 2,3,4,5-tetrachlorophenol (2,3,4,5-TTCP); 2,3,4,6-tetrachlorophenol (2,3,4,6-TTCP); and pentachlorophenol (PCP). Using this method Morgade et al. [296] found levels of pentachlorophenol between less than 0.14mg L-1 and 340µg L-1 in Florida potable water and 0.14mg L-1 and 0.1µg L-1 in well water. No other chlorophenols were found.
Chlorophenols
415
5.8.2.2 Gas chromatography–mass spectrometry Sithole et al. [297] used selected ion gas chromatography–mass spectrometry to determine halogenated phenols in raw and potable water. They compared two derivatization procedures, in situ acetylation and the formation of pentafluorobenzyl derivative for determining chlorophenols and bromophenols in raw and potable water. Some problems which prevented the use of the pentafluorobenzyl derivative are discussed. Using in situ acetylation, gas chromatography with electron-capture detection was compared with gas chromatography–mass spectrometry with selective ion monitoring. The sensitivity of electroncapture detection could not be fully exploited because of background response. The selective ion monitoring technique was less susceptible to interference and more sensitive to phenol and monohalogenated phenols. Recoveries of phenols from water samples were generally more than 80%. Treated water samples showed a variety of halogenated phenols which were not present in the raw water indicating these compounds had been formed in the treatment process. 5.8.2.3 High-performance liquid chromatography This technique has been applied to the determination of pentachlorophenol in potable water [298]. 5.8.3 Waste waters 5.8.3.1 Gas chromatography Chau and Coburn [299] have described an electron-capture gas chromatographic method for the determination of pentachlorophenol in natural waste waters. The phenol is extracted from the sample (1L) into benzene and subsequently from the latter into 0.1M potassium carbonate. Addition of acetic anhydride to the aqueous solution gives the acetate derivative, which is extracted into hexane and analysed by gas chromatography with the use of a conventional polar column. The limit of detection of 10pg L-1 of sample. Matthew and Elzerman [300] directly acetylated substituted phenols and determined them by gas chromatography of methylene dichloride extracts. Acetylation was achieved by addition of acetic anhydride to an alkaline aqueous solution of phenols. The results of the analysis of four synthetic mixtures of six phenols spiked into distilled water (at 0.8– 40µg L-1) are given. For dye waste waters, direct acetylation accomplished a considerable, although not complete, elimination of coextractives. The
416
Halogen-containing compounds
application of direct acetylation to municipal waste waters is considered promising as they contain fewer coextractives. Buisson et al. [301] determined nanogram per litre levels of chlorinated phenols in waste water by capillary gas chromatography with electron-capture detection. Derivatization, by alkylation with pentafluorobenzoyl chloride enhanced the response of the electroncapture detector, gave stable derivatives, improved selectivity and volatility, reduced background and reagent peaks and provided a simple methodology. Spiked samples showed a mean recovery of 70% and a relative standard deviation of 7–15%. The American Public Health Authority Standard Method series includes gas chromatographic methods for the determination in waste waters of pentachlorophenol. To determine pentachlorophenol, the sample is acidified (pH3) and pentachlorophenol extracted with an organic solvent, methylated with diazomethane and analysed by electron-capture gas chromatography. Further identification of pentachlorophenol is made with a mass spectrometer. 5.8.3.2 High-performance liquid chromatography Column chromatographic techniques have been described [285] for the determination of pentachlorophenol and other chlorophenols, but these methods are not suitable for the determination of low levels. The best method for determining pentachlorophenol is conversion into the methyl ether followed by analysis using gas chromatography with an electroncapture detector, or gas chromatography coupled with mass spectrometry. Both of these methods require an extensive amount of pretreatment and highly trained personnel for the operation of the equipment. Ervin and McGinnis [285] attempted to overcome this problem by developing a high-performance liquid chromatographic method for determining in water low concentrations of pentachlorophenol and chlorinated impurities that occur in the technical grade material such as 2,3,4,6-tetrachlorophenol, mono-, di- and trichlorophenols, octaheptaand hexachlorodibenzo-p-dioxins, and a variety of other polychlorinated aromatic compounds. The method involves chloroform extraction of acidified waste water samples and rotary evaporation without heat. After redissolving in chloroform the samples were analysed directly by high-performance liquid chromatography on a microparticulate silica gel column. A number of solvent combinations are possible and cyclohexane–acetic acid (92:2v/v) is preferred. The minimum detectable concentration is
Halocarboxylic acids
417
1mg L-1 (without sample concentration) and the coefficient of variation is 1–2%. Melcher et al. [302] used an on-line liquid chromatography analyser for the determination of down to 3µg L -1 of pentachlorophenol, dichlorophenol and tetrachlorophenol in waste waters. 5.8.3.3 Preconcentration Rostad et al. [303] used a cyclohexyl bonded phase resin column to isolate creosote and pentachlorophenol in ground water samples taken at a waste site. 5.8.4 Trade effluents 5.8.4.1 Gas chromatography–mass spectrometry Gas chromatography–mass spectrometry has been applied to the identification and determination of chlorophenols in spent bleach liquor trade effluents from sulphate plants [304, 305]. 5.8.5 Sewage effluents 5.8.5.1 Gas chromatography–mass spectrometry Gas chromatography–mass spectrometry has been used to measure the occurrence of pentachlorophenol and hexachlorophene in water and sewage [306]. 5.8.5.2 Miscellaneous Baird et al. [307] utilized gas chromatography in their study of the biodegradability of chlorinated phenols in sewage sludge. 5.9 Halocarboxylic acids 5.9.1 Natural waters 5.9.1.1 Gas chromatography Ozawa and Tsukioka [308] have described a gas chromatographic method for determining down to 0.6µg L-1 of sodium monofluoroacetate in natural waters.
418
Halogen-containing compounds
5.9.1.2 Isotope dilution mass spectrometry This technique has been used to determine trichloroacetic acid in natural waters [309, 310]. 5.9.1.3 Miscellaneous Trifluoroacetic acid is a significant breakdown product of chlorofluorocarbons present in the atmosphere and water and is therefore a marker for the presence of these substances in the ecosystem. Zehavi and Seiber [333] have discussed a headspace, gas chromatographic method for the determination of microgram per litre levels of trifluoroacetic acid in fog and rain water and surface waters. The described method determines trace levels of trifluoroacetic acid, an atmospheric breakdown product of several of the hydrofluorocarbon and hydrochlorofluorocarbon replacements for the chlorofluorocarbon refrigerants in water and air. Trifluoroacetic acid is derivatized to the volatile methyl trifluoroacetate and determined by automated headspace gas chromatography (HSGC) with electron-capture detection or manual HSGC using gas chromatography–mass spectrometry in the selected ion monitoring (SIM) mode. The method is based on the reaction of an aqueous sample containing trifluoroacetic acid with dimethyl sulphate in concentrated sulphuric acid in a sealed headspace vial under conditions favouring distribution of methyltrifluoroacetate to the vapour phase. Water samples are prepared by evaporative concentration, during which trifluoroacetic acid is retained as the anion, followed by extraction with diethyl ether of the acidified sample and then back-extraction of trifluoroacetic acid (as the anion) in aqueous bicarbonate solution. The extraction step is required for samples with a relatively high background of other salts and organic materials. Air samples are collected in sodium bicarbonate–glycerincoated glass denuder tubes and prepared by rinsing the denuder contents with water to form an aqueous sample for derivatization and analysis. Recoveries of trifluoroacetic acid from spiked water, with and without evaporative concentration, and from spiked air were quantitative, with estimated detection limits of 10ng mL -1 (unconcentrated) and 25pg mL-1 (concentrated 250mL:1mL) for water. Several environmental air, fog water, rain water and surface water samples were successfully analysed; many showed the presence of trifluoroacetic acid.
Halocarboxylic acids
419
5.9.2 Potable waters 5.9.2.1 Pyrolysis method Stachel et al. [311] determined halogenated organic compounds in potable water using mineralization at high temperature in a stream of oxygen. The procedure is carried out in a pyrolysis tube in which the sample is injected into a stream of carbon dioxide carrier gas. The carrier gas is passed through a superheated steam zone at 700°C and a pre-oxidation and cracking zone at 1100°C, in a stream of oxygen. The sample, carrier gas and oxygen are mixed in a pyrolysis chamber at 850°C and passed through a second 850°C heating zone before cooling and titration. The mineralization achieved was about 100% for polychlorinated biphenyl and hexachlorobenzol and 91% for dichloroacetic acid. Uden and Miller [312] reported a qualitative and quantitative investigation of some non-volatile chlorinated organics found in chlorinated potable waters. Concentrations of di- and trichloroacetic acids were in the range 30–160µg L-1, while those of chloral were lower by an order of magnitude. The concentrations of these compounds were readily measurable using a microwave-emission plasma detector. All compounds identified were also formed during the laboratory chlorination of soil fulvic acids. 5.9.3 Trade effluents 5.9.3.1 Isotachophoresis Onodera et al. [313] examined the applicability of isotachophoresis to the identification and determination of chlorinated mono- and dicarboxylic acids in chlorinated effluents. Four electrolyte systems for the separation of the acids were evaluated. The potential unit values in each system were determined for the chlorinated acids. A mechanism for the reaction of phenol with hypochlorite in dilute aqueous solutions is suggested, based on results from the isotachophoretic analysis of diethyl ether extracts taken from phenol treated with hypochlorite. 5.9.4 Sewage effluents 5.9.4.1 Miscellaneous Chlorophenoxyisobutyric acid has been determined [314] in primary treated sewage works effluent and river waters.
420
Halogen-containing compounds
5.10 Chlorinated isocyanurates 5.10.1 Swimming pool water 5.10.1.1 Spectrophotometric method These substances have been determined in amounts down to 10mg L1 in swimming pool water by a spectrophotometric procedure [315]. The method is based on the production of isocyanuric acid and reaction with cobalt acetate, in the presence of isopropylamine. The water sample is evaporated and the residue dissolved in methanol; 5mL is mixed with 5mL chloroform. Cobalt acetate solution is added followed by 0.5mL isopropylamine. After 30min, the extinction is read at 565nm against a blank of methanol–chloroform–isopropylamine (10:10:1). 5.11 Chlorolignosulphonic acids 5.11.1 Natural waters 5.11.1.1 Pyrolysis-gas chromatography Van Loon et al. [316] used a technique combining pyrolysis–gas chromatography and mass spectrometry with single-ion monitoring to determine macromolecular (mol wt > 1000) Chlorolignosulphonic acids in natural waters in amounts down to 0.1–0.2µg L-1. 5.11.2 Potable waters 5.11.2.1 Pyrolysis–gas chromatography The pyrolysis–gas chromatography method [316] discussed in Section 5.11.1.1 has been used to analyse potable waters. 5.11.3 Trade effluents 5.11.3.1 Pyrolysis–gas chromatography The pyrolysis–gas chromatography method [316] described in Section 5.11.1.1 has been applied to mill effluents.
Polychlorodibenzo-p-dioxins and polychlorodibenzofurans
421
5.12 Polychlorodibenzo-p-dioxins and polychlorodibenzofurans 5.12.1 Natural waters 5.12.1.1 Gas chromatography–mass spectrometry Yasuhara and Itoh [317] have described a gas chromatographic method combined with mass spectrometric detection for the identification and determination of 2,3,7,8-tetrachlorodibenzo-p-dioxin in river water. Taguchi et al. [318] applied high-resolution mass spectrometry coupled with selective ion monitoring (HRMS-SIM) to the determination of polychlorodibenzo-p-dioxins, and tetra-, octa-, chloroand polychlorodibenzofurans in natural waters. Laramee et al. [319] applied negative-ion mass spectrometry to polychlorodibenzo-p-dioxins and found a correlation between observed mass fragmentations and calculated internal energies. Fifteen cogeners and isomers were examined. Schimmel et al. [320] studied the molar response of polychlorodibenzo-p-dioxins, and polychlorodibenzofurans and polychlorobiphenyls using a mass spectrometric detector. Plomley et al. [321] studied the mass spectrometry of polychlorodibenzo-pdioxins and polychlorodibenzofurans in a quadrupole ion trap and compared results obtained in single-frequency modulation and multifrequency resonant excitation modes. These workers concluded that small-bandwidth multifrequency irradiation (MFI) represents the best mode of resonant excitation for the mass spectrometry–mass spectrometry analysis of multiple target analytes such as polychlorodibenzo-p-dioxins and polychlorodibenzofurans. 5.12.1.2 High-performance liquid chromatography Thielen and Olsen [322] optimized conditions for the extraction from water on alumina of tetrachlorodibenzo-p-dioxins and their isomers and carried out a specific determination of 2,3,7,8-tetrachlorodibenzo-p-dioxin. Alumina separates dioxins and polychlorobiphenyls. Dioxins can be desorbed from alumina with 12% chloroform in benzene. Further isolation of the 2,3,7,8-dioxin isomer from others is made with either 15% methylene dichloride, 40% chloroform or with 100% n-butyl chloride. Gonzalez et al. [331] used reversed-phase liquid chromatography to determine polychlorodibenzofurans in natural waters. Kimata et al. [323] used electron-acceptor and electron-donor highperformance liquid chromatographic stationary phases for the reversedphase liquid chromatographic separation and identification of polychlorodibenzo-p-dioxins.
422
Halogen-containing compounds
5.12.2 Potable waters 5.12.2.1 Gas chromatography O’Keefe et al. [324] used gas chromatography to determine down to 1pg L1 of 2,3,7,8-tetrachlorodibenzo-p-dioxin in potable water. 5.12.3 Waste waters 5.12.3.1 Miscellaneous Peters et al. [325] have discussed the determination of 2,3,7,8tetrachlorodibenzo-p-dioxin in waste water. 5.13 Chloroanisoles and chloromethylanisoles 5.13.1 Natural waters 5.13.1.1 Gas chromatography–mass spectrometry This technique [326] has been used to determine down to 0.02µg L-1 of these substances in natural waters. 5.14 Polybromobiphenyls 5.14.1 Natural waters 5.14.1.1 Irradiation–gas chromatography Polybrominated biphenyls have been detected in environmental samples by a procedure involving ultraviolet irradiation followed by gas chromatography of the photodegradation products [327–330]. 5.15 Polychloroterphenyls 5.15.1 Natural waters 5.15.1.1 Gas chromatography–mass spectrometry The occurrence of these substances has been reported in river water [327]. The samples were extracted with hexane. The extracts were cleaned by
References
423
column chromatography, on either alumina or Florasil. The analysis of polychlorinated terphenyls was carried out with a combination of a mass spectrometer and a gas chromatograph used in the mass fragmentography mode. Two m/e values were selected (m/e = 436 and m/ e = 470) and from all samples mass fragmentograms were taken on these m/e values. The gas chromatographic conditions were: 3ft × 1/8 in o.d. (0.9m × 3mm) Pyrex column, filled 3% OV-17 on Varaport 30. Temperature programming was used: 6°C min-1, 200–285°C for m/e = 436 and 8°C min-1, 200–285°C for m/e = 470; 4°C min-1, 125–200°C for m/e = 256; 6°C min-1, 125–200°C for m/e = 290; and 6°C min-1, 150–250°C for m/e = 324. Approximately 0.1ppm of polychlorinated terphenyl was found in river water; the corresponding figure for PCBs was 0.1ppb. References 1 Murray, A.J. and Riley, J.P. Anal. Chim. Acta 65 261 (1973) 2 Murray, A.J. and Riley, J.P. Nature (London) 242, 37 (1973) 3 Novak, J., Zluticky, J., Kubulka, V. and Mostecky, J. Journal of Chromatography 76 45, (1973) 4 Kleopfer, R.D. and Fairless, B.T. Environ. Sci. Technol. 6 1036 (1972) 5 Jensen, S., Jernelov, A., Large, R. and Palmork, K.H. Food and Agriculture Organisation of the United Nations. Report No. FIR: MP 170/E-88 November (1970) 6 Dietz, F. and Traud, J. Vom Water 41 137 (1973) 7 Zitko, V. J. Chromat. 81 152 (1973) 8 Zitko, V. J. Ass. Off. Anal. Chem. 57 1253 (1974) 9 Friedman, D. and Lombardo, P. J. Ass. Off. Anal. Chem. 58 703 (1975) 10 Deetman, A.A., Demeulemeester, P., Garcia, M., Hauck, G., Hollies, J.I., Krockenberger, D., Palin, D.E., Prigge, H., Rohrschneider, L. and Schmidthammer, L. Anal. Chim. Acta 82 1 (1976) 11 Guam, C.S. and Wong, M.K. Journal of Chromatography 72 283 (1972) 12 Hagenmaier, H., Werner, G. and Jager, W. Z. Wasser Abwasser Forsch. 15 195 (1982) 13 Chiba, K. and Haraguchi, H. Analytical Chemistry 55 1504 (1983) 14 Kaiser, K.L.E. and Oliver, B.G. Analytical Chemistry 48 2207 (1976) 15 Hrivnak, F., Siskupic, P. and Hassler, J. Vodni Hospodarstivi. Series B 28 195 (1978) 16 Hellman, H. Z. Wasser Abwasser Forsch. 21 67 (1988) 17 McLary, T.A. and Barker, J.F. Groundwater Monit. Rev. 7 63 (1988) 18 Biebier Dammann, V., Funk, W., Von Mareard, G. and Rinne, D. Z. Wasser Abwasser Forsch. 20 22 (1987) 19 Yurteri, C, Ryan, D.F., Callow, J.J. and Gurol, M.D., J. Water Pollut. Contr. Fed. 59 950 (1987) 20 Hellman, H. Z. Wasser Abwasser Forsch. 18 92 (1985) 21 Mehran, M., Cooper, M. and Jennings, W. Journal of Chromatographic Science 24 142 (1986) 22 Kirschen, N. American Laboratory 16 60 (1984) 23 Prath, K.F. and Pauliszyn, J. Analytical Chemistry 64 2101 (1992)
424
Halogen-containing compounds
24 Lopez-Avila, V., Heath, N. and Hu, A.J. Journal of Chromatographic Science 25 351 (1987) 25 Cochran, J.W. J. High Resol. Chromatog. 10 573 (1987) 26 Chichester-Constable, D.J., Barbeau, M.E., Liu, S.L., Smith, S.R. and Stuart, J.D. Anal. Lett. (London) 20 403 (1987) 27 Mosesman, N.H., Sidisky, L.M. and Corman, S.D. Journal of Chromatographic Science 25 351 (1987) 28 Hollies, J.I., Pinnington, P.F., Handley, A.J., Baldwin, M.K. and Bennett, D. Anal. Chim. Acta 111 201 (1979) 29 Kummert, R., Molnar-Kbuica, E. and Giger, W. Analytical Chemistry 50 1637 (1978) 30 Stozek, A. and Beumer, W. Korrespondenz Abswasser 26 632 (1979) 32 Simonov, V.D., Popova, L.N. and Shamsutdinov, T.M. Dokl. Neftehim. Sekt. Basgkirsk. Resp. Provl. Vses. Khim. Obshch. Mendeleeva 6, 357 (1971) Ref. Zh. Khim. 19GD Abstract No. 59302 (1972) 33 Dilling, W.L., Tefertiller, O. and Kallos, G.J. Environ. Sci. Technol. 9 833 (1975) 34 Hellman, H. Z. Wasser Abwasser Forsch. 18 210 (1985) 35 Friedman, L.C., Schroder, J. and Brooks, M.G. Environ. Sci. Technol. 20 826 (1986) 36 Martinsen, K., Kringstad, A. and Carlberg, G.E. Water Sci. Technol. 20 13 (1988) 37 Pankow, J.F. J. High Resol. Chromat. Chromat. Commun. 6 292 (1983) 38 Dawson R., Diley, J.P. and Tennant, R.H. Marine Chemistry 4 83 (1976) 39 Glaze, W., Henderson, J.E., Bell, J.E. and Van Wheeler, A. Journal of Chromatographic Science 11 580 (1973) 40 Lukacovic, L., Mikulas, M., Vanko, A. and Kiss, G. Journal of Chromatography 207 373 (1981) 41 Pfannhauser, W. and Thaller, A. Fresenius Z. Analyt. Chemie, 322 220 (1985) 42 American Public Health Association, Standard Methods for the Examination of Waters and Waste Waters. Method for Volatile Chlorinated Organic Compounds in Water and Waste Waters (1985) 43 Von Duzzeln, J., Lahl, U., Stachel, B., and Thiemann, W. Z. Wasser Abwasser Forsch. 15 272 (1982) 44 Henderson, J.E. and Glaze, W.H. Water Research 16 211 (1982) 45 H.M. Stationery Office, London. Methods for the Examination of Waters and Associated Materials (1986). Chlorobenzene in Water Organochlorine Pesticides and PCB’s in Turbid Waters, Halogenated Solvents and Related Compounds in Sewage Sludge (1985) 46 Rudolph, K.U. and Koppke, K.E. Korrespond. Abwasser 35 990 (1988) 47 Hassler, J. and Rippa, F. Vyskummy Ustai Vodneko Hospodarstra; Veda a Vyskum Praxi. No. 50 50 pp (1977) 48 Eklund, G., Josefsson, B. and Roos, C. J. High Resol. Chromat. Chromat. Commun. 1 34 (1978) 49 Eklund, G., Josefsson, B. and Roos, C. Journal of Chromatography 142 575 (1977) 50 Albert, S. and Jonke, B. Zeitschrift für Wasser und Abwasser 8 140 (1975) 51 Burgasser, A.J. and Calaruotolo, J.F. Analytical Chemistry 49 1508 (1977) 52 Narang, R.S. and Bush, B. Analytical Chemistry 52 2076 (1980) 53 Fujii, T. Analytical Chemistry 49 1985 (1977) 54 Rivera, J., Cuberes, M.R. and Albaiges, J. Bull. Environ. Contamin. Toxicol. 18 624 (1977)
References
425
55 Stein, U.B. and Narang, R.S. Bulletin of Environmental Contamination and Toxicology 27 583 (1981) 56 Bellar, T.A. Lichtenberg, J.J. and Eichelberger, J.W. Environ. Sci. Technol. 10 926 (1976) 57 Benoit, G.M. and Williams, D.T. Bull. Environmental Contamination and Toxicology 27 303 (1981) 58 Ando, M. and Sayato, Y. Water Research 18 315 (1984) 59 Determination of Vinyl Monomer in aqueous effluents, Analytical Chemistry Branch, Southeast Environmental Research Laboratory, Environmental Protection Agency , Athens, Ga, USA (1974) 60 Okumura, K., Kawuda, J. and Uno, T. Analyst ( London ) 107 1498 (1982) 61 Quimby, B.R., Delaney, M.F., Uden, P.C. and Barnes, R.M. Analytical Chemistry 52 259 (1980) 62 Kroner, R.C. Public Works 111 81 (1980) 63 Libbey, A.J. Analyst ( London ) 111 1221 (1986) 64 Keith, L.H. Identification and Analyses of Organic Pollutants in Water, Ann Arbor Science Publishers Inc., Ann Arbor, MI (1976) 65 Brodtmann, N.V. J. Am. Water Works Ass. 67 558 (1975) 66 Coburn, J.A., Valdmanis, I.A. and Chau, A.S.Y. The Extraction of Organochlorine Pesticides and P.C.B.’s from Natural Waters with XAD-2, Water Quality Branch, Canada Centre of Inland Waters, Burlington, Ontario, L7R 4A6 Canada 67 Musty, P.R. and Nickless, G.J. Chromat. 89 185 (1974) 68 Junk, G.A., Rickard, J.J., Grieser, M.D., Witlak, D., Witlak, J.L., Arguello, M.D., Vick, R., Svec H.J., Fritz, V.S. and Calder, G.V. Journal of Chromatography 99 745 (1974) 69 Coburn, J.A., Valdmanis, I.A. and Chau, A.S.U. J. Ass. Off. Anal Chem. 60 224 (1977) 70 Kissinger, L.D. and Fritz, J.S. J. Am. Water Works Ass. 68 435 (1976) 71 Junk, G.A., Chriswell, D.C., Chiang, R.C., Kissinger, L.D., Richard, J.J., Fritz, J.S. and Svec, H.J. Fresenius Z. Anal. Chem. 282 331 (1976) 72 Arbon, R.E. and Grimsrud, E.P. Analytical Chemistry 62 1762 (1990) 73 Bognar, J.A., Knighton, W.B. and Grimsrud, E.P. Analytical Chemistry 64 2451 (1992) 74 Symons, J.M., Bell, T.A., Carswell, J.K., DeMarco, J., Kropp, K.L., Robeck, G.G., Seegar, D.R., Slocum, D.J., Smith, B.L. and Stevens, A.A. J. Am. Water Works Ass. 67 643 (1975) 75 Keith, L.H. Identification and Analysis of Organic Pollutants in Water, Ann Arbor Science Publishers, Inc., Ann Arbor, ML (1976) 76 National Survey for Halomethanes in Drinking Water, Health and Welfare Canada, 77-EHD-9 (1977) 77 Bush, B. and Narang, R.S. Syrotyn Bulletin of Environmental Science and Technology 13 436 (1977) 78 Morris, R.L. and Johnson, L.G. J. Amer. Water Works Association 68 492 (1976) 79 Friant, S.L. Thesis, Drexel University, University Microfilms Ltd. London, 468 pr 29162 (1972) 80 Gomella, C. and Belle, J.P. Techniques et Sciences Municipales 73 125 (1978) 81 Varma, M.M., Siddique, M.R., Doty, K.T. and Machis, H. J. Amer. Water Works Association 71 389 (1979) 82 Dietz, E.A. and Singley, K.F. Analytical Chemistry 51 1809 (1979) 83 Rook, J.J. Water Treat. Exam. 21 259 (1972) 84 Rook, J.J. Water Treat. Exam. 23 234 (1974)
426
85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106
107 108 109 110 111 112 113 114 115
Halogen-containing compounds
Lovelock, J.E. Nature ( London ) 256 193 (1975) Lovelock, J.E., Maggo, R.J. and Wade, R.J. Nature ( London ) 241 194 (1973) Zlatkis, A., Lichtenstein, A. and Tuchbee, A. Chromatographia 8 67 (1973) Mieure, J.P. and Dietrich, M.M. J. Chromatog. Sci. 11 559 (1973) Dowty, B. and Laseter, J.L. Analytical Letters 8 25 (1975) Bellar, T.A. and Lichtenberg, J.J. J. Am. Water Works Ass. 66 739 (1974) Grob, K. Journal of Chromatography 84 255 (1973) Grob, K. and Grob, G. Journal of Chromatography 90 303 (1974) Grob, K., Grob, K. Jnr. and Grob, G. Journal of Chromatography 106 299 (1975) Bertsch, W., Andersson, E. and Holzer, G. Journal of Chromatography 112 701 (1975) Stevens, A.A. and Symons, J.M. Environmental Impact of Water Chlorination. Oak Ridge National Lab., Oak Ridge, Tennessee (1975) Dressman, R.C., Stevens, A.A., Fair, J. and Smith, B. J. Amer. Water Works Assoc. 71 392 (1979) US Environmental Protection Agency. Part II I. Appendix C. Analysis of Trihalomethanes in Drinking Water. Federal Register, 44, No 231 to 68672. Nov. 29th (1979) Method 501.1 The Analysis of Trihalomethanes in Finished Waters by the Purge and Trap Method. US Environmental Protection Agency, EMSL, Cincinnati, Ohio, 45268. Nov 6th (1979) Anonymous. Chemical Engineering News 54 35 (1976) Ligon, V.V. and Johnson, R.I. Analytical Chemistry 48 481 (1976) Brass, H.J. Journal of American Laboratory 12 23 (1980) Kirschen, N.A. Varian Instrument Application 15 2 (1981) Method 60 US Environmental Protection Agency, Federal Register 69464, 44, No 223, Dec 3rd (1979) Packard Technical News 1 No 7258 , February (1981) Pierce, C.T., Grochowski, R.J., Kongovi, R.R., Narangajvanaca, K. and Brock, G.L. American Laboratory 34, April (1981) Department of the Environment/National Water Council Standing Committee of Analysts. H.M. Stationery Office, London. Methods for the Examination of Waters and Associated Materials. Chloro and Bromo-Tri Halogenated Methanes in Water (1981) Leppine, L. and Archambault, J.F. Analytical Chemistry 64 810 (1992) Janda, V. Vodni Hospodarstvi Series B 31 137 (1981) Schnoor, J.L., Nitzschke, J.L., Lucas, R.D. and Veenstra, J.N. Environ. Sci. Technol, 113 1134 (1979) Dretsman, R.C. and Stevens, A.A. J. American Water Works Association 75 431 (1983) Oake, R.J. and Anderson, I.M. Water Research Centre, Medmenham, UK Report No. TR 217. The determination of carbon adsorbable organohalide in water (1984) Blanchard, R.D. and Hardy, J.K. Analytical Chemistry 50 1621 (1984) Tardiff, R.G. and Dunzer, M. Toxicity of Organic Compounds in Drinking Water, Environmental Protection Agency, Cincinnati, OH (1974) Report on the Carcinogenesic Assay of Chloroform, US National Cancer Institute Bethesda, MD (1976) Cantor, R.H., Mason, T.J. and McCabe, L.J. Association of cancer mortality rates and trihalomethane levels in municipal drinking water supplies. Proc. 10th Ann. Mtg. Soc. Epidemio. Res. (1977)
References
427
116 Analytical Report – New Orleans Area Water Supply Study EPA 906/ 10-74 -002. Surveillance and Analysis Division, USEOA Region V1, Dallas, Texas. Mimeo, 30 (1974) 117 The Implications of Cancer-Causing Substances in Mississippi River Water, Environmental Defence Fund, Washington DC (1974) 118 Glaze, W.H. and Rawley, R. J. Am. Water Works Ass. 71 509 (1979) 119 Luong, T., Peters, C.J., Young, R.J. and Perry, R. Environ. Technol Lett. 1 299 (1980) 120 Bunn, W.W., Haas, B.B., Deane, E.R. and Kleopfer, R.D. Environ. Lett. 10 205 (1975) 121 Glaze, W.H., Henderson, J.E. and Smith, G. Private communication 122 Dowty, B.K., Carlisle, D.R. and Laseter, J.L. Environ. Sci Technol. 9 762 (1975) 123 New Orleans Area Water Supply Study (Draft Analytical Report), Lower Mississippi River Facility, USEPA (1974) 124 Stevens, A.A., Slocum, C.J., Seiger, D.R. and Robeck, G.G. Cong. Environ. Impact of Water Chlorination, Oak Ridge National Laboratory, Oak Ridge, TN (1975) 125 Symons, J.M., Bellar, T.A., Carswell, J.K., Demarco, J., Kropp, K.L., Roebeck, G.G., Seegar, D.R., Slocum, C.J., Smith, B.L. and Stevens, A.A. National Organics Reconnaissance Survey for Halogenated Organics in Drinking Water, EPA, Cincinnati, OH (1975) 126 Tardiff, R.G. and Dunzer, M. Toxicity of Organic Compounds in Drinking Water, EPA, Cincinnati, OH (1973) 127 Tabor, M.W. Environ. Sci. Technol. 17 324 (1983) 128 US Environmental Protection Agency Federal Register No 28, 43 5756 (1978) 129 USEPA Control of Organic Chemical Contaminants in Drinking Water, Federation Register, 44 5755 et seq. (Feb 9, 1978) 130 Harris, L.E., Budde, W.L. and Eichelberger, J.W. Analyst ( London ) 46 1912 (1974) 131 Nicolson, A.A. and Meresz, O. Bull. Environ. Contam. Toxicol. 14 4 (1975) 132 Nicolson, A.A. and Meresz, O. Bull. Environ. Contam. Toxicol 14 453 (1975) 133 Fielding, M., McLoughlin, K. and Steel, C. Water Research Centre Enquiry Report ER 532, August 1977. Water Research Centre, Stevenage Laboratory, Elden Way, Stevenage, Herts, UK (1977) 134 Fam, S. and Stenstrom, M.K. J. Water Pollut. Contr. Fed. 59 969 (1987) 135 De Leer, E.W.B. In Aqueous Chlorination Products the Origin of Oranochlorine Compounds in Drinking and Surface Waters, Delft University Press (1987) 136 Smillie, R.D., Nicolson, A.A., Meresz, O. Duholke, W.K., Rees, G.A.V., Roberts, K. and Fung, C. Organics in Ontario Drinking Waters, Part II. A Survey of Selected Water Treatment Plants, Ontario Ministry of the Environment, April 1977 137 Richard, J.J. and Junk, G.A. J. Am. Water Works Ass. 69 62 (1977) 138 Mieure, J.P. J. Am. Water Works Ass. 69 62 (1977) 139 Nicolson, A.A., Meresz, O. and Lemyk, B. Analytical Chemistry 49 814 (1977) 140 Hammerstrand, K. Chloroform in Drinking Water, Varian Instrument Application, 10 22 (1976) 141 Viryasov, M.B. and Nikitin, Y.S. Soviet Journal of Water Chemistry and Technology 9 71 (1987) 142 Eklund, G., Josefsson, B. and Roos, C.J. High Resolution Chromatography. Chromat Commun. 1 34 (1978) 143 Dingynan, H. and Jianfei, T. Analytical Chemistry 63 2078 (1991) 144 Goldberg, M.C., Delong, L. and Sinclair, M. Anal. Chem. 45 89 (1973)
428
Halogen-containing compounds
145 Henderson, J.E., Peyton, G.R. and Glaze, W.H. Recommended procedure: a convenient liquid–liquid extraction method for the determination of chloroform and other halomethanes in water. Rpt. Trace Analysis Lab. Inst. Applied Sci., North Texas State Univ., Denton, TX (1976) 146 Webb, R.G. Isolating Organic Water Pollutants: XAD Resins, Urethane Foams, Solvent Extraction. USEPA-660/4-75 003 pp. 17–18 (1975) 147 Trussell, A.R. Microextraction of Organic Pollutants: Application to Trihalomethanes in Drinking Water. President’s Fellowship Pt. Univ. of California, San Diego (1976) 148 Reichert, J.K. and Lochtman, J. Environmental Science Letters 4 15 (1983) 149 Von Rensburg, J.F.T., Van Huyssteen, J.T. and Hassett, A.J. Water Research 12 127 (1978) 150 Method 501.2, The Analysis of Trihalomethanes in Drinking Water by Liquid/Liquid Extraction, USEPA, EMSL, Cincinnati, OH 45268 (November 6, 1979) 151 Henderson, J.E., Peyton, G.R. and Glaze, W.H. A convenient liquid/liquid extraction method for the determination of halomethane in water at the parts per billion level. Identification and Analysis of Organic Pollutants in Water (L.H. Keith, Editor) Ann Arbor Science, Ann Arbor, MI, 105, 195 (1976) 152 Symons, J.M. National Organics Reconnaissance Survey for Halogenated Organics in Drinking Water, J. A. Water Works Ass. 11 634 (1975) 153 Bellar, T.A. and Lichtenberg, J.J. J. Am. Water Works Ass. 12 739 (1974) 154 Kirschen, N.A. Varian Instrument Applications 14 10 (1980) 155 Lange, A.L. and Kawczynski, E. J. Am. Water Works Ass. 70 653 (1978) 156 Norin, H. and Renberg, L. Water Research 14 1397 (1980) 157 Henderson, J.E., Peyton, G.R. and Glaze, W.H. A convenient liquid–liquid extraction method for the determination of halomethane in water at the parts per billion level. Identification and Analysis of Organic Pollutants in Water (L.H. Keith, Editor). Ann Arbor Science, Ann Arbor, Michigan, 105, 195 (1976) 158 Mehran, M.F., Slither, R.A. and Cooper W.J. J. Chromatographic Science 22 241 (1984) 159 Inoko, M., Tsuchiya, M. and Matsumo, T. Environmental Bulletin (Series B) 7 129 (1984) 160 Trussell, A.R., Umphres, M.D., Leong, L.Y.C. and Trussell, R.R. J. Amer. Water Works Assoc. 71 385 (1979) 161 Reding, R.J. Chromatographic Science 25 338 (1987) 162 Kissinger, L.D. US National Technical Information Service, Springfield, VA. Report No I5-T-845 161 pp. (32411) (1979) 163 McCarthy, L.V., Overton, E.B., Raschke, C.K. and Laseter, J.L. Anal Lett. 13 1417 (1980) 164 Comba, M.E. and Kaiser, K.L.E. International Journal of Environmental Analytical Chemistry 16 17 (1983) 165 Peruzi, P. and Griffini, O. Journal of High Resolution Chromatography, Chromatography Communications 8 450 (1985) 166 Nicolson, B.C., Bursill, D.B. and Conche, D.J. Journal of Chromatography 325 221 (1985) 167 Marshall, J.W. and Wampler, T.P. American Laboratory 20 72 (1988) 168 Fujii, T. Journal of Chromatography 139 297 (1977) 169 Fujii, T. Anal. Chim. Acta 92 117 (1977) 170 Fujii, T. Bulletin of the Chemical Society of Japan 50 2911 (1977)
References
429
171 Vogel, T.M. and Reinhard, M. Environ. Sci. Technol 20 992 (1986) 172 Coleman, W.E., Munch, J.W., Kayler, W.H., Streicher, R.P., Rinhand, H.P. and Meier, J. Environ. Sci. Technol. 18 674 (1984) 173 Christman, R.F., Norwood, D.L., Webb, M.R. and Bobenrieth, M.J. US National Technical Information Service, Springfield, VA, Report No. PB81161952. Chlorination of aquatic humic substances (1981) 174 Pruden, A.L. and Ollis, D.F. Environmental Science and Technology 17 628 (1983) 175 Morris, R.L. and Johnson, L.G. J. Amer. Water Works Ass. 68 492 (1976) 176 Suffet, L.H. and Radziul, O. J. Amer. Water Works Assoc. 68 520 (1976) 177 Montiel, A. Tribune de Labedeau 32 422 (1979) 178 Simmonds, P.G. and Kerns E. Journal of Chromatography 186 785 (1979) 179 Otson, R., William, D.T. and Bothwell, P.D. Environ. Sci. Technol. 13 936 (1979) 180 Castello, G., Gerhino, T.C. and Kanitz, S. Journal of Chromatography 247 263 (1982) 181 Castello, G., Gerbino, T.C. and Kanitz, S. J. Chromat. 351 165 (1986) 182 Croll, B.T., Summer, M.E. and Leathard, D.A. Analyst (London) 111 73 (1986) 183 Chilli, L. and Borgioli, A. Water Research 20 273 (1986) 184 Quimby, B.D. and Delaney, M.F. Analytical Chemistry 51 875 (1979) 185 Quimby, B.D., Uden, P.C. and Barnes, P.C. Analytical Chemistry 50 2112 (1978) 186 Pfaender, F.E., Jones, B.B., Stevens, A.M., Moore, L. and Hass, J.R. Environmental Science and Technology 12 438 (1978) 187 US Federal Register. 69464 44 No 223 Dec 3rd (1979) 188 Chiba, K. and Haraguchi, H. Analytical Chemistry 55 1504 (1983) 189 Wong, T. and Lenahan, B. Bulletin of Environmental Contamination and Toxicology 32 429 (1984) 190 Watts, C.D. Water Research Centre Technical Report TR 110. Mass spectrometric identification of non-volatile organic compounds (1979) 191 Reunanen, M. and Kronfeld, R. Journal of Chromatographic Science 20 449 (1982) 192 Janda, V., Marha, K. and Mitera, J. Journal of High Resolution Chromatography, Chromatography Communications 11 54 (1988) 193 Miller, J.W. and Uden, P.C. Environmental Science and Technology 17 150 (1983) 194 Hahn, J. Korrespond Abwasser 29 539 (1952) 195 Heckel, E. Umwelt, No 1 31 (1986) 196 De Leer, E.W.B. H O 13 171 (1980) 197 Weil, L., Jandik, J.2 and Eicheldorfer, D. Z. Wasser Abwasser Forsch. 13 141 (1980) 198 Kapila, S. and Aue, W.A. Journal of Chromatography 15 569 (1977) 199 Oliver, B.G. and Bothen, K.D. Analytical Chemistry 52 2066 (1980) 200 Italia, M.P. and Uden, P.C. Journal of Chromatography 449 326 (1988) 201 Ryan, D.A., Argentine, S.M. and Rice, G.W. Analytical Chemistry 62 853 (1990) 202 Di Corcia, A. and Samperi, R. Analytical Chemistry 62 1490 (1990) 203 Stetter, J.R. and Cao, Z. Analytical Chemistry 62 1490 (1990) 204 Bjorseth, A., Carlberg, G.E. and Moller, M. Science of the Total Environment 11 197 (1979) 205 Oyler, A.R., Bodenner, D.L., Welch, K.J., Hukhaxen, R.K., Carlsen, R.M., Kopperman, H.L. and Caple, R.L. Analytical Chemistry 50 837 (1978)
430
Halogen-containing compounds
206 Webb, R.G. and McCall A.C. J. Ass. Off. Anal. Chem. 55 746 (1972) 207 Bauer, U. Gas- u. Wasserfach. Wasser Abwasser 113 58 (1972) 208 Berg, O.W., Diolady, P.L. and Rees, G.A. Bulletin of Environmental Contamination and Toxicology 7 338 (1972) 209 Beezhold, F.L. and Stout, V.F. Bulletin of Environmental Contamination and Toxicology 10 10 (1973) 210 Chromopak News. June 1979 No. 20. Chromopak Ltd. PO Box 3, 4330 AA Middelburg, The Netherlands (1979) 211 Schulte, E. and Acker, L. Z. Analyt. Chem. 268 260 (1974) 212 Brinkmann, A.Th., Seetz, J.W.F.L. and Reymer, H.G.M. Journal of Chromatography 116 353 (1976) 213 Brinkmann, A.Th., DeKok, A., De Vries, G. and Reymer, H.G.M. Journal of Chromatography 128 101 (1976) 214 Dunn, W.J., Stalling, D.L., Schwartz, T.R., Hogan, J.W., Petty, J.D., Johonsson, E. and Wold, S. Analytical Chemistry 56 1308 (1984) 215 Tas, A.C. and Kleipool, R.J.C. Bulletin of Environmental Contamination and Toxicology 8 32 (1972) 216 Sissons, D. and Welti, D. Journal of Chromatography 60 15 (1971) 217 Albro, P.W., Haseman, J.K., Clemmer, T. and Corbett, B.J. Journal of Chromatography 136 147 (1977) 218 Sawyer, L.D. J. Ass. Off. Analytical Chemists 61 272 (1978) 219 Albro, P.W. and Fishbein, O. Journal of Chromatography 69 273 (1972) 220 Hanai, T. and Walton, J.F. Analytical Chemistry 49 1954 (1977) 221 Sawhrey, B.L., Frink, C.R. and Glowa, W. Journal of Environmental Quality 10 444 (1981) 222 Duinker, J.C., Schulz, D.E. and Petrick, G. Marine Pollution Bulletin 19 19 (1988) 223 Hussun, M.N. and Juns, P.C. Analytical Chemistry 60 978 (1988) 224 Roblat, A., Xyra-fas, G. and Marshall, D. Analytical Chemistry 60 982 (1988) 225 Stalling, D.L. and Huckins, J.N. J. Ass. Off. Anal. Chem. 54 801 (1972) 226 Ahnoff, M. and Josefsson, B. Anal. Lett. 6 1083 (1973) 227 Webb, R.G. and McCall, A.C. J. Chromat.Sci. 11 366 (1973) 228 Eichelberger, J.W., Harris, L.E. and Budde, W.J. Analytical Chemistry 46 227 (1974) 229 Karlruber, B.A., Hormann, W.D. and Ramsteinen, K.A. Analytical Chemistry 47 2453 (1975) 230 Ahnoff, M. and Josefsson, B. Anal. Lett. 6 1036 (1973) 231 Voyksner, R.D., Hass, J.R. and Sovocoal, G.W. Analytical Chemistry 55 744 (1983) 232 Pellazari, E.E., Moseley, M.A. and Cooper, S.D. Journal of Chromatography 334 277 (1985) 233 Tanabe, S., Kannan, N., Wakimoto, T. and Tatsukawa, R. International Journal of Environmental Analytical Chemistry 29 199 (1987) 234 de Vos, R.H. and Peet, E.W. Bulletin of Environmental Contamination and Toxicology 6 164 (1971) 235 Sackmauer, O.M., Pal’usova, O. and Szokolay, A. Water Research 11 551 (1977) 236 Kaminsky, L.S. and Fasko, M.J. Journal of Chromatography 155 363 (1978) 237 Needham, L.L., Surek, A.L., Head, S.L., Burse, V.W. and Liddle, J.A. Analytical Chemistry 52 2227 (1980) 238 Dong, M.N. and DiCesare, J.L. Journal of Chromatographic Science 20 517 (1982)
References
431
239 Petrick, G., Schulz, D.E. and Duinker, J.C. Journal of Chromatography 435 241 (1988) 240 Farwell, S.O., Beland, F.A. and Geer, R.D. Bull. Environ. Contam. Toxicol. 10 157 (1973) 241 Neely, W.B. Science of the Total Environment 7 117 (1977) 242 Kopanica, M. Science of the Total Environment 37 83 (1984) 243 Aga, D.S., Thurman, E.M. and Dowes, M.L. Analytical Chemistry 66 1495 (1994) 244 Gesser, H.D., Chow, A., Davis, F.C., Uthe, J.F and Reinke, J. Anal Lett. 4 883 (1971) 245 Uthe, J.F., Reinke, J. and Gesser, H. Environ. Lett. 3 117 (1972) 246 Ahnoff, M. and Josefsson, B. Bulletin of Environmental Contamination and Toxicology 13 159 (1975) 247 US Environmental Protection Agency. US National Information Service, Springfield, VA . Report No. PB 279547 (1977) (30381) 248 Inland Waters Directorate. Analytical Methods Manual. Water Quality Branch, Ottawa, Ontario, Part 2, Organic Constituents (1974) 249 Coburn, J.A., Valdmanis, I.A. and Chau, A.S.Y. J. Ass. Off. Anal. Chem. 60 224 (1977) 250 Chriswell, C.D., Ericson, R.L., Junk, G.A., Lee, K.W., Fritz, J.S. and Svec, H.J. J. Amer Water Works Assoc. 69 669 (1977) 251 Musty, P.R. and Nickless, G. Journal of Chromatography 100 83 (1974) 252 Musty, P.R. and Nickless, G. Journal of Chromatography 120 369 (1976) 253 Gesser, H.D., Chow, A., Davis, F.C., Uthe, J.F and Reinke, J. Anal. Lett. 4 883 (1971) 254 Bedford, J.W. Bull. Environ. Contam. Toxicol. 12 662 (1974) 255 Lawrence, J. and Todine, H.M. Environmental Science and Technology 10 381 (1976) 256 Mantoura, R.F.C. and Lewellyn, C.A. Anal. Chim. Acta 151 297 (1983) 257 Thome, J.P. and Vandaele, Y. International Journal of Environmental Analytical Chemistry 29 95 (1987) 258 Elder, D. Marine Pollution Bulletin 7 63 (1976) 259 Sullivan, K.F, Atlas, E.L. and Glain, C.S. Analytical Chemistry 53 1718 (1981) 260 Le’Bel, G.L. and Williams, D.T. Bull. Environ. Contam. Toxicol. 24 397 (1980) 261 Le’Bel, G.L., Williams, D.T., Griffith, G. and Benoit, F.M. J. Ass. Off. Anal. Chem. 62 281 (1979) 262 Delfino, J.J. and Easty, N. Anal. Chem. 51 2235 (1979) 263 Easty, D.B. and Wabers, B.A. Tappi 61 71 (1978) 264 Easty, D.B. and Wabers, B.A. Anal. Lett. 10 857 (1977) 265 Lopez-Avila, V., Schoen, S., Milanes, J. and Beckert, W.F. J. Assoc. Off. Anal. Chem. 71 375 (1988) 266 Canagay, A.B. and Levine, P.L. US Environmental Protection Agency. Cincinnati, Ohio , Report EPA-600/ 2-79 -166, 100 pp. (P22CAR) (1979) 267 Skinner, R.F., Fins, W.F. and Banelli, E.J. Finnigan Spectra 3 No. 1 (1973) 268 Herbst, E., Schennert, I., Kein, W. and Korte, F. Chemosphere 6 725 (1977) 269 Furukawa, K., Tonomura, K. and Kawtayashi, A. Applied Environmental Microbiology 35 223 (1978) 270 Gaffney, P.E. Journal of Water Pollution Control 49 401 (1977) 271 Carr, R.C., Thomas, P. and Nett, J.M. Bull. Environ. Contam. Technol. 28 477 (1982) 272 Stark, A. Journal of Agricultural and Food Chemistry 17 871 (1969) 273 Rudling, L. Water Research 4 533 (1970)
432
274 275 276 277 278 279 280 281 282 283 284 285 286 287 288 289 290 291 292 293 294 295 296 297 298 299 300 301 302 303 304 305 306 307 308
Halogen-containing compounds
Renberg, L. Analytical Chemistry 46 459 (1974) Chriswell, C. and Cheng, R. Analytical Chemistry 47 1325 (1975) Farrington, D.S. and Mundy, W. Analyst (London) 101 639 (1976) Hoben, H.J., Ching, S.A., Casarett, L.J. and Young, R.A. Bulletin of Environmental Contamination and Toxicology 15 78 (1976) Ashiya, K., Otani, H. and Kajino, K. Japan Water Works Association No. 536 12 (1979) Korhonen, I.O.O. and Knuutinen, J. Chromatographia 17 154 (1983) Bengtsson, G. Journal of Chromatographic Science 23 397 (1985) Henshaw, B.G., Morgan, J.W.W. and Williams, N. Journal of Chromatography 110 37 (1975) Zigler, M.G. and Phillips, W.F. Environ. Sci. Technol. 1 65 (1967) Wolkoff, A.W. and Larose, R.H. J. Chromat. 99 731 (1974) Thielemann, H. and Luther, M. Pharmazie 25 367 (1970) Ervin, H.E. and McGinnis, G.D. Journal of Chromatography 190 203 (1980) Brandt, G. and Kettrup, A. Fresenius Z. Analyst Chemie 327 213 (1987) Wade, A.L., Hankridge, F.M. and Williams, A.P. Anal. Chim. Acta 105 91 (1979) Boyle, T.P., Robinson–Wilson, E.F., Petty, J.D. and Weber, W. Bulletin of Environmental Contamination and Toxicology 24 178 (1980) Ingram, L.I., McGinnis, G.D. and Poriki, S.V. Analytical Chemistry 51 1077 (1979) Malissa, H., Szolyenyi, G. and Winsauer, K. Fresenius Z. Analyst Chemie 321 17 (1985) Noren, K. and Sjovall, J. Journal of Chromatography 414 55 (1987) Gaivdzik, J., Gaivdzik, B. and Czerwinska-Billi, A. Chromatographia 25 504 (1988) Noll, K.E. and Gounaris, A. Water Research 22 815 (1988) Fingler, S., Drevenkar, V. and Vasitic, Z. Microchim. Acta No 4/6 163 (1987) Chriswell, C. and Cheng, R. Analytical Chemistry 47 1325 (1975) Morgade, C., Barquet, A. and Pfaffenherger, C.D. Bulletin of Environmental Contamination and Toxicology 24 257 (1980) Sithole, B.B., Williams, D.T., Lastoria, C. and Robertson J.L. Journal Ass. Off. Anal. Chem. 69 466 (1986) McMurtrey, K.D., Holcomb, A.E., Ekwenchi, A.U. and Fawcett, N.C. Journal of Liquid Chromatography 7 953 (1984) Chau, A.S.Y. and Coburn, J.A. Journal Ass. Off. Anal. Chem. 57 389 (1974) Matthew, J. and Elzerman, A.W. Analytical Letters (London) 14 1351 (1981) Buisson, R.S.K., Kirk, P.W.W. and Lester, J.N. Journal of Chromatographic Science 22 339 (1984) Melcher, R.G., Dakker, D.W. and Hughes G.H. Analytical Chemistry 64 2258 (1992) Rostad, C.E., Perrira, W.E. and Ratcliff, S.M. Analytical Chemistry 56 2856 (1984) Lindstrom, K. and Nordin, J. Journal of Chromatography 128 13 (1976) Carlberg, G.E., Gjos, N., Maller, M., Gustavsen, K.O., Tveten, G. and Renberg, L. Sci. Total Environ. 15 3 (1980) Buhler, D.R., Rasmussan, M.E. and Nakaue, H.S. Environmental Science and Technology 7 929 (1973) Baird, R.B., Kuo, C.L., Shapiro, J.S. and Yonko, W.A. Archives of Environmental Contamination and Toxicology 2 165 (1974) Ozawa, H. and Tsukioka, T. Analytical Chemistry 59 2914 (1987)
References
433
309 Norwood, D.L., Christman, R.F., Johnson, J.D. and Hass J.R. Journal American Water Works Assoc. 78 175 (1986) 310 Miller, J.W., Uden, P.C. and Barnes, R.M. Analytical Chemistry 54 485 (1982) 311 Stachel, B.. Lahl, U., Kozicki, R., Pobbulski, A. and Thiemann, W. Chemosphere 11 803 (1982) 312 Uden, P.C. and Miller, J.W. Journal Amer. Water Works Association 75 524 (1983) 313 Onodera, S., Udagawa, T., Tabata, M., Ishkura, S. and Susuki, S. Journal of Chromatography 287 176 (1984) 314 Hignite, C. and Azarnoff, D.L. Life Science 20 337 (1977) 315 Van der Haar, G., Pijper-Noordhoft, F.M. and Strikwenda, K. H O 12 240 2 (1972) 316 Van Loon, W.H.G.M., Bron, J.S. and de Groot, B. Analytical Chemistry 65 1726 (1993) 317 Yasuhara, A. and Itoh, M.M. Environmental Science and Technology 21 971 (1987) 318 Taguchi, V.V., Reiner, E.J., Wong, D.T., Meresz, D. and Hassas, B. Analytical Chemistry 60 1429 (1988) 319 Laramee, J.A., Arbogast, B.C. and Dienzer, M.L. Analytical Chemistry 60 1937 (1988) 320 Schimmel, H., Schmid, B., Backer, R. and Ballschmitter, K. Analytical Chemistry 65 640 (1993) 321 Plomley, J.B., March, R.E. and Mercer, R.S. Analytical Chemistry 68 2345 (1996) 322 Thielen, D.R. and Olsen, G. Analytical Chemistry 60 1332 (1988) 323 Kimata, K., Hosoya, K., Araki, T., Tanaka, N., Barnhard, E.R., Alexander, L.R., Sirimanne, S., McLlure, P.C., Granger, J. and Patterson, P.C. Analytical Chemistry 65 2502 (1993) 324 O’Keefe, P., Meyer, C., Smith, R., Hilker, D., Aldous, K. and Wilson, L. Chemosphere 15 1127 (1986) 325 Peters, T.L., Nestrick, T.J. and Lamparski, L.L. Water Research 18 1021 (1984) 326 Lee, H.P. Journal Ass. Off. Anal. Chem. 71 803 (1988) 327 Freudenthal, J. and Greve, P.A. Bull. Environ. Contam. Toxicol. 10 108 (1973) 328 Banks, K.A. and Bills, D.D. Journal of Chromatography 33 450 (1968) 329 Potter, W.G. LIB 1609 , FDA, Minneapolis District (1969) 330 Erney, D.R. J. Am. Oil Colour Ass. 58 1202 (1975) 331 Gonzalez, E.B., Baumann, R.A., Gooijer, C., Velthorst, W.H. and Frei, R.U. Chemosphere 16 1123 (1987) 332 Kaiser, L.L.E. and Wong, P.T.S. Bulletin of Environmental Contamination and Toxicology 11 291 (1974) 333 Zehavi, D. and Seiber, J.N. Analytical Chemistry 68 3450 (1996) 334 Burguera, L. and Burguera, J.L. Anal. Chim. Acta 153 53 (1983)
Chapter 6
Nitrogen-containing compounds
6.1 Aliphatic amines 6.1.1 Natural waters 6.1.1.1 Spectrophotometric methods Florence and Farner [1] determined parts per billion (109) amines in waters, sea water and raffinates from uranium processing using a spectrophotometric procedure. 6.1.1.2 Gas chromatography Hermanson et al. [2] used an aluminium column (276cm × 4mm) packed with 80–100 mesh Chromosorb W supporting 8.9% of amine 220 at 95°C with nitrogen as carrier gas and flame ionization detection. A rectilinear response was obtained between peak area and amount of propylamine, dipropylamine and propanol; between 0.2 and 2.0µg. Gas flame ionization chromatography has been used to determine dimethylamine [3, 4], dimethylformamide [3], propylamine [2] and diisopropylamine [2] in river water and industrial effluents. To separate C –C mono-, di- and trialkylamines, Onuska [4] adjusted the pH of the 1 4 sample to between 5 and 8. A lµL aliquot of the filtrate was injected on to a stainless steel column (185cm × 2mm i.d.) packed with 28% of Pennwalt 223 and 4% of potassium hydroxide on Gas-Chrom R (80–100 mesh) and maintained at 134°C. A dual-flame ionization detector was used and the carrier gas was helium (flow rate 52.2mL min-1). The detector response was rectilinear between 10ng and at least 100µg of dimethylamine, and the reproducibility was good. The column could be regenerated by increasing the column temperature to greater than 180°C. Kimoto et al. [5] used gas chromatography to determine simple aliphatic amines in dichloromethane extracts of water samples. They
Aliphatic amines
435
used a gas chromatograph equipped with a nitrogen–phosphorus detector and a 1.8m × 2mm i.d. glass column packed with Carbopack B– Carbowax 20-potassium hydroxide. The flow rates were helium 20mL min-1 and air 80mL min-1. The injector and detector temperatures were 200 and 250°C respectively. The column was programmed from 70 to 150°C at 4°C min-1 and held at the upper temperature for an additional 4min. Then 2µL samples were injected. 6.1.1.3 Gas chromatography–mass spectrometry Avery and Junk [6] converted trace levels of primary aliphatic amines in aqueous solutions to imine derivatives using pentafluorobenzaldehyde. The derivatives were extracted into hexane measured by combined highresolution gas chromatography–mass spectrometry using multiple-ion monitoring. The method was successfully applied to the analysis of primary amines in potable water and river water down to a detection limit of 10µg L-1 for 0.5mL samples. 6.1.1.4 Thin-layer chromatography Hexane 1,6-diamine has been determined in amounts down to 0.5µg L-1 in water by thin-layer chromatography and paper chromatography [7]. 6.1.1.5 Liquid chromatography Nishikawa [8] described a liquid chromatographic procedure for the determination of aliphatic diamines as their acetylacetenates in natural water with a detection limit of 1.4–20µg L-1. 6.1.2 Sea water 6.1.2.1 Spectrophotometric methods Florence and Farner [1] determined parts per million of aliphatic amines in sea water using Spectrophotometric procedures. 6.1.2.2 Miscellaneous Yang et al. [9] determined nanomolar quantities of individual low molecular weight amines (and organic acids) in sea water. Amines were diffused from the sample across a hydrophilic membrane to concentrate and separate them from inorganic salts and
436
Nitrogen-containing compounds
most other dissolved organic compounds. Methylamine, dimethylamine and trimethylamine were all found in measurable amounts in sea water. Petty et al. [177] used flow injection sample processing with fluorescence detection for the determination of total primary amines in sea water. The effects of carrier stream flow rate and dispersion tube length on sensitivity and sampling rates were studied. Relative selective responses of several amino acids and other primary amines were determined using two dispersion tube lengths. Linear calibration curves were obtained over the ranges 0–10-6M and 0–10-8M glycine. Precisions of better than 2% at 10-6M and a detection limit of 10-8M glycine were obtained. 6.1.3 Potable waters 6.1.3.1 Gas chromatography–mass spectrometry The gas chromatographic–mass spectrometric method [6] described in Section 6.1.1.3 has been applied to the determination of down to 10µg L-1 of primary aliphatic amines in potable water. 6.1.4 Trade effluents 6.1.4.1 Titration procedure Fishman and Pevzner [10] determined down to 0.15mg L-1 of hexamine in effluents from polyformaldehyde manufacture by potentiometric titration with 0.2M hydrochloric acid. 6.1.4.2 Spectrophotometric method Florence and Farner [1] determined parts per million of aliphatic amines in raffinates from uranium processing using Spectrophotometric procedures. 6.1.4.3 Gas chromatography The gas chromatographic procedures [2–4] described in Section 6.1.1.2 have been used to determine dimethylamine, dimethylformamide, propylamine and diisopropylamine in industrial effluents. 6.1.4.4 Thin-layer chromatography Hexane 1,6-diamine has been determined in amounts down to 5µg L-1 in water and industrial effluents by thin-layer chromatography and paper chromatography [11].
Aromatic amines
437
6.1.5 High-purity waters 6.1.5.1 Thin-layer chromatography Octadecylamine, a corrosion-inhibiting boiler water additive, has been isolated from water by solvent extraction with ethylene dichloride or by trapping on a micro Chromosorb column [12]. The levels of octadecylamine were determined using high-performance thin-layer chromatography and reflectance densitometry. Recovery at 3mg L-1 averaged 81.3% using solvent extraction and 94.0% with the column. Recovery at 0.3mg L-1 was 94.2% using the column procedure. Other permitted boiler water additives did not interfere with the analysis. Sherma and Pallasta [13] determined cyclohexylamine in water by solidphase extraction and high-performance thin-layer chromatography. 6.2 Aromatic amines 6.2.1 Natural waters 6.2.1.1 Spectrophotometric method Aniline derivatives at the microgram level have been determined in natural waters by a Spectrophotometric procedure involving diazotization with sulphuric acid and naphthol [14]. 6.2.1.2 Gas chromatography Wegman and DeKorte [15] have described a method for the determination of aniline, toluidines, monochloroanilines and their derivatives in surface water. The aromatic amines in an iso-octane extract of the water sample were brominated and determined by gas chromatography with a capillary column and an electron-capture detector. The results of analysis of Rhine water samples in the Netherlands was presented. The detection limits were in the 5–15ng L-1 range. 6.2.1.3 High-performance liquid chromatography Benzidine has been determined in natural water using column chromatography [16]. 6.2.1.4 Miscellaneous Stuber and Leenheer [17] assessed the aqueous elution, selective concentration approach for isolating aromatic bases from water, and the
438
Nitrogen-containing compounds
factors controlling the concentration process identified using polymethacrylic ester as an adsorbent. The degree of concentration attainable depends on the ratio of the capacity of the natural form of the amine to that of the ionized form. The capacity factors of ionic forms of amines on polymethylacrylic ester resins are 20–250 times lower than those of the neutral forms and increase with the hydrophobicity of the amine. 6.2.1.5 Preconcentration Nielen et al. [18] used a strongly acidic cation-exchange resin for on-line preconcentration of polar anilines in water. The method could be automated and a detection limit for the nine anilines examined corresponding to 0.02 to 0.5µg L-1 was obtained in river water samples. 6.2.2 Sea water 6.2.2.1 High-performance liquid chromatography Varney and Preston [19] discussed the measurement of trace aromatic amines in estuary and sea water using high-performance liquid chromatography. Aniline, methyl aniline, 1-naphthylamine and diphenylamine at trace levels were determined using this technique and electrochemical detection. Two electrochemical detectors (a thinlayer, dual glassy carbon electrode cell and a dual porous electrode system) were compared. The electrochemical behaviour of the compounds was investigated using hydrodynamic and cyclic voltammetry. Detection limits of 15 and 1.5nM were achieved using coulometric and amperometric cells respectively when using an in-line preconcentration step. 6.2.3 Trade effluents 6.2.3.1 Spectrophotometric method Aniline derivatives at the microgram level have been determined in natural waters by a Spectrophotometric procedure involving diazotization with sulphuric acid and naphthol [14]. 6.2.3.2 Gas chromatography Diphenylamine and other aromatic bases have been determined respectively in industrial effluents [20] and coal carbonization products [21] utilizing gas chromatography.
Chloroaromatic amines
439
6.2.3.3 Miscellaneous Jenkins [22] determined benzidine at the microgram per litre concentration range in effluents by spectrophotometric, gas chromatographic and thin-layer chromatographic procedures. 6.2.4 Waste waters 6.2.4.1 Spectrophotometric method The American Public Health Association [23] has described a method for the determination of benzidine and its salts at the 0.3µg L-1 level in water and waste water. In this method the water sample is made basic and the benzidine is extracted with ethyl acetate. Clean-up is accomplished by extracting the benzidine from the ethyl acetate with hydrochloric acid. Chloramine-T is added to the acid solution to oxidize the benzidine. The yellow oxidization product is extracted with ethyl acetate and measured with a scanning spectrophotometer. The spectrum from 510nm to 370nm is used for qualitative identification. 6.2.4.2 Gas chromatography Aniline and its derivatives have been determined [24] in waste water at the low microgram per litre level by gas chromatography with thermionic nitrogen–phosphorus detection. Precision is generally 5– 15% relative standard deviation and recoveries are generally 75% or better. 6.2.4.3 Miscellaneous Jenkins [22] determined benzidine at the nanogram per litre concentration range in waste waters by spectrophotometric, gas chromatographic and thin-layer chromatographic procedures. 6.3 Chloroaromatic amines 6.3.1 Natural waters 6.3.1.1 Liquid chromatography Di Corcia and Samperi [25] have described a method for the determination of down to 0.1µg L-1 of chloroaniline in environmental waters. Extraction of the sample with acetonitrile with two traps in tandem is followed by liquid chromatographic analysis.
440
Nitrogen-containing compounds
6.3.2 Waste waters 6.3.2.1 Gas chromatography Trace levels of chloroanilines and chloronitroanilines have been determined in waterworks sludges using a technique based on gas chromatography using simultaneously Hall and NP-specific detectors [26]. 6.4 Aminophenols 6.4.1 Natural waters 6.4.1.1 Gas chromatography A gas chromatographic procedure has been described for the determination of down to 0.1mol of aminophenols in natural waters [27]. 6.4.2 Potable waters 6.4.2.1 High-performance liquid chromatography High-performance liquid chromatography has been applied to the determination of 4-aminophenol in potable water [28]. 6.5 Nitro compounds 6.5.1 Natural waters 6.5.1.1 Spectrophotometric method Heller et al. [29] have described a Spectrophotometric method for the determination of trinitrotoluene in natural waters. 6.5.1.2 High-performance liquid chromatography This technique has been applied to the determination of nitro compounds in natural waters [30]. 6.5.2 Potable waters 6.5.2.1 High-performance liquid chromatography This technique has been used to determine traces of ethylene glycol dinitrate in potable water [31].
Nitro compounds
441
6.5.2.2 Gas chromatography Patil [32] determined nitrobenzine and 2,4-dinitrophenol in potable water by gas chromatography. 6.5.3 Trade effluents 6.5.3.1 Spectrophotometric methods Trinitrotoluene has been determined in effluents from munitions plants by Spectrophotometric [33, 34] and fluorimetric procedures [35]. The fluorimetric procedure described by Heller et al. [35] is capable of determining down to 0.07µg L -1 trinitrotoluene in munitions plant wastes. The basis of the method is the absorption of trinitrotoluene on a quaternary ammonium ion-exchange resin which darkens on contact with trinitrotoluene. The resin is irradiated with ultraviolet radiation and the fluorescent output is monitored by a photomultiplier. Total aliphatic and aromatic nitro compounds have been determined in industrial effluents in amounts down to l.5mg L-1 by a method [36] based on the chemical reduction of all nitro compounds in the sample to amino compounds using titanium chloride in hydrochloric acid as reducing agent. For the final Spectrophotometric determination the amines are reacted with sodium naphthoquinone sulphonate. The method is sensitive, reliable and precise. 6.5.3.2 Gas chromatography Richard and Junk [37] use solid phase extraction using XAD-4 resin elution with ethyl acetate and separation by gas chromatography to determine microgram per litre quantities of nitro compounds in munitions works effluents. A method for the quantitative analysis of explosive compounds in effluents has been described [38] which includes 2,4-TNT, teryl, 2,4-DNT and 2,6-DNT at the microgram per litre level. The compounds were extracted from water with toluene and analysed by capillary gas chromatography with electron-capture detection. Recoveries of 96– 103% were obtained in samples spiked at 1, 10 and 100µg L-1. The stabilities of these compounds in water at pH values ranging from 0.5 to 10 were studied over 33 days. The compounds were stable at pH1–4 and significant losses occurred at pH values greater than 6. 6.5.3.3 Gas chromatography–mass spectrometry Spanggord et al. [39] used gas chromatography–mass spectrometry to identify over 30 nitroaromatic compounds present in effluents
442
Nitrogen-containing compounds
produced during the manufacture of trinitrotoluene. Three major components were consistently identified and represented about 75% of the total component distribution, 2,4- and 2,6-dinitrotoluene and 1,3dinitrobenzene. 6.5.3.4 Preconcentration A method has been described [40] for the preconcentration of nitro compounds in munitions works effluents by adsorption on Poropak resins. Following desorption with acetone the nitro compounds (nitramines, nitrotoluenes and nitroaliphatics) were measured by highperformance liquid chromatography at a gold–mercury electrode. The results of analysis of munitions in test samples showed that Poropak resins were superior in performance to RDX resins. Analysis time was approximately 2h per sample for isolation and quantification. Detection limits of 1µg L-1 were approached. 6.5.4 Waste waters 6.5.4.1 High-performance liquid chromatography This technique has been used to determine nitro compounds in munition plant wastes [41]. Godjehann et al. [168] have applied on-line high-performance liquid chromatography–proton NMR to the determination of nitroaromatic compounds in ammunition plant ground waters. This is the first time these two techniques have been coupled. Using a continuous flow mode at very low sample flow rates of about 0.017mL m-1 and large injection volumes of 400µL the confirmation of the identity of many different types of nitroaromatic compounds was achieved at the microgram per litre level after solid-phase extraction of the ground water sample at a flow rate of 0.006µL m-1. Less than 29nmol (5µg) of 1:3 dinitrobenzene was determined when injected on to a 75mm × 4mm reversed phase C column. 18
6.5.4.2 Miscellaneous Spangoord and Suta [42] characterized various nitrocompounds in waste waters produced during the manufacture of trinitrotoluene. Heller et al. [35] have described a semiquantitative method, using fluorescent ion-exchange resins for the monitoring of trinitrotoluene in munition plant waste water.
Nitrophenols
443
6.6 Nitrophenols 6.6.1 Natural waters 6.6.1.1 Gas chromatography Cranmer [43] has described a gas chromatographic method for the determination of down to 0.05ppm 4-nitrophenol in human urine and this procedure would, no doubt, be applicable to water samples. The sample is adjusted to pH11 and twice extracted with benzene–ethyl ether (4:1); the extracts are discarded, the aqueous phase is acidified to pH2 and the extraction is repeated. This extract is dried over sodium sulphate and hexamethyldisilazane is added. To effect on-column silanization, the solution is injected into a gas chromatograph equipped with an inlet operated at 200°C, an electron-capture detector and a column of either 5% of DC-200 on Gas-Chrom Q or 1.5% of OV-17 plus 1.95% of QF-1 on Chrom W, operated at 140°C. Bengtsson [44] has described optional conditions for the determination of trace amounts of nitrophenols (0.01–0.2µg L-1) and dinitrophenols (1.6µg L-1). The phenols are toluene extracted from the water sample, derivativized with heptafluorobutyricanhydride and separated and quantified by electron-capture gas chromatography or flame ionization gas chromatography. 6.6.1.2 Thin-layer chromatography Chambou and Chambon [45] have described a thin-layer chromatographic method for the determination of down to 5µg L -1 of 4,6-dinitro-o-cresol in diethyl ether extracts of water samples. The extract is applied to a 0.25mm layer of Cellulose MN and the chromatogram is developed with ammonia-saturated butanol (in a chamber equilibrated with the solvent) for 14cm. The nitrophenol spots are located by the bright yellow colour formed with ammonia, removed from the plate and extracted with 50% acetic acid (0.05mL)–methanol (5mL); the extinction of the extract is measured at 370nm [47]. 6.6.1.3 Column chromatography This technique coupled with a spectrophotometric detection has been used to determine mixtures of nitrophenols in natural waters [46].
444
Nitrogen-containing compounds
6.6.1.4 Polarography Micro amounts down to 4µg L-1 of dinitro-o-cresol have been determined in water by oscillographic polarography. 6.7 Nitriles 6.7.1 Natural waters 6.7.1.1 Titration and spectrophotometric methods Stefanescu and Ursu [48] determined acrylonitrile and acetonitrile in residual waters by spectrophotometric and titrimetric procedures after separation from the sample by azeotropic distillation. Acrylonitrile and acetonitrile are determined together (as ammonia) after alkyline hydrolysis and acrylonitrile is determined separately with mercaptoacetic acid. Down to 2mg of each substance per litre can be determined. Ghersin et al. [49] compared colorimetric and titrimetric procedures for the determination of acrylonitrile in waters. A titration method based on addition of sodium sulphide to the acrylonitrile followed by titration of liberated sodium hydroxide gave a sensitivity of 20mg L-1 acrylonitrile. Provided that the sample is not yellow or brown in colour, a titrimetric method involving the use of mercaptoacetic acid has a sensitivity of 2mg acrylonitrile per litre, or down to 0.4mg L-1 if the acrylonitrile is separated by a preliminary distillation from methanol sulphuric acid medium; such a distillation also separates acrylonitrile from interfering substances. 6.7.1.2 Gas chromatography This technique has been used to determine acrylonitrile at the microgram per litre level in natural waters [50]. 6.8 Amides 6.8.1 Natural waters 6.8.1.1 Gas chromatography Arkell and Croll [51] and Croll and Simpkins [52] determined acrylamide monomer in amounts down to 0.1µg L-1 by a gas chromatographic procedure.
Amides
445
The gas chromatographic procedure [2–4] described in Section 6.1.1.2 has been used to determine dimethyl formamide in natural waters. Hashimoto [53] used gas chromatography to determine acrylamide monomer in water. This method utilizes the response of the brominated form of acrylamide (a,ß-dibromopropionamide) to electron-capture detection. The acrylamide is brominated by an ionic reaction using bromine in the presence of potassium bromide and hydrobromic acid. The a,ß-dibromopropionamide thus formed is extracted from water using ethyl acetate and the concentrate is then analysed by electroncapture gas–liquid chromatography. Electron-capture detectors are, however, prone to contamination. Difficulties in maintaining optimum electron-capture sensitivity over extended periods may lead to some decrease in sensitivity during analysis. 6.8.1.2 High-performance liquid chromatography Earlier high-performance liquid chromatographic methods [54, 55] for the determination of acrylamide monomer had detection limits of approximately 0.1µg L-1. Brown and Rhead [56] improved the sensitivity of high-performance liquid chromatography to 0.2µg L-1. The procedure consists of bromination, extraction of the a,ß-dibromopropionamide with ethyl acetate and quantification using high-performance liquid chromatography with ultraviolet detection. Samples tested included river, sea and estuarine waters, sewage and china clay works effluents, and potable waters. The levels of inorganic ultraviolet absorbing impurities found in water samples did not interfere in this procedure. The solvent extraction procedure lowered interferences in all samples tested without removal of acrylamide or excessive use of solvents. The experimental yields of a,ß-dibromopropionamide encountered gave a mean of 70.13 ± 8.52% (95% confidence level) for acrylamide-spiked waters, over the concentration range 0.2-8.0µg L -1 of acrylamide monomer. 6.8.1.3 Polarography This technique has been used to determine acrylamide in natural waters [57]. 6.8.2 Sea water 6.8.2.1 High-performance liquid chromatography The technique [56] discussed in Section 6.8.1.2 has been applied to sea and estuarine waters.
446
Nitrogen-containing compounds
6.8.3 Potable waters 6.8.3.1 Titration method A titration procedure [58] has been described for the determination of amides in potable waters. 6.8.3.2 Spectrophotometric method Cationic polyelectrolytes have been determined in water [59] using a method based on changes in light absorbance resulting from the formation of an insoluble change complex with ponceau 5 dye. 6.8.3.3 High-performance liquid chromatography Acrylamide has been determined in potable water by high performance liquid chromatography with ultraviolet detection [60]. The procedure [56] described in Section 6.8.1.2 has been applied to the determination of an acrylonitrile in potable water. 6.8.4 Trade effluents 6.8.4.1 Flow injection analysis Ludi et al. [61] carried out an enzymic determination of dimethylformamide in process effluents using flow injection analysis. 6.8.4.2 Gas chromatography The gas chromatographic procedures described in Section 6.1.1.2 [2–4] have been used to determine dimethylformamide in industrial effluents. 6.8.4.3 High-performance liquid chromatography This technique has been applied to the determination of acrylamide in trade effluents [62]. The procedure described by Brown and Rhead [56] and discussed in Section 6.8.1.2 has also been applied to trade effluents. 6.8.4.4 Size exclusion chromatography Leung et al. [63] determined polyacrylamides in coal washery effluents by ultrafiltration size exclusion chromatography with ultraviolet detection. A column of TSK 5000 PW hydrophilic and semi-rigid porous polymer gel was used with 0.05M sodium sulphate as mobile phase.
Nitrosamines
447
6.8.4.5 Miscellaneous Jungreis [64] has described a simple technique for the microdetermination of polymer flocculants in processed mine water which involves measuring the laminar flow rate of the mine water through a membrane filter under vacuum. Bezazyan et al. [65] have described a phototurbidimetric method for the determination of polyacrylamide flocculants in effluents. 6.8.5 Sewage effluents 6.8.5.1 High-performance liquid chromatography The technique described by Brown and Rhead [56] and discussed in Section 6.8.1.2 has been applied to the determination of acrylamide in sewage effluents. 6.9 Nitrosamines Many N-nitrosamines are toxic and carcinogenic, and furthermore the carcinogenic action exhibits a high degree of organ specificity. Nitrosamines are formed by interaction between nitrate and an amine with varying ease, depending on the nature of the amine and the prevailing conditions. The reaction is not restricted to secondary amines, but also occurs with primary and tertiary amines and even quaternary ammonium salts. Thus, the precursors are widespread, both as naturally occurring compounds and as generated in many commercial and industrial processes, and it is therefore conceivable that trace amounts may be present in air and water in the vicinity of industrial sites. Nitrosamines in minute amounts have been found in deionized water generated from the resins. 6.9.1 Natural waters Mills and Alexander [66] have discussed the factors affecting the formation of dimethylnitrosamine in samples of water and soil. Dimethylnitrosamine was formed as readily in sterilized samples as in non-sterile samples, indicating that, although microorganisms can carry an enzymatic nitrosation in some soils and waters, dimethylnitrosamine can be formed by a non-enzymatic reaction, even if near neutral conditions. The presence of organic matter appears to be important in promoting nitrosation in the presence of the requisite precursors. Various workers have discussed the determination of nitrosamines in natural waters [66–68].
448
Nitrogen-containing compounds
6.9.1.1 Gas chromatography Nikaido et al. [67] give details of procedures for the recovery of low levels of dialkylnitrosamines from natural waters, including lake water and sewage effluents before subsequent detection by gas chromatography. The recovery technique involves the addition of potassium carbonate to the sample and concentration of the nitroso compounds on Amberlite XAD-2 resin. Greater than 90% recoveries were obtained for dimethylnitrosamine and diethylnitrosamine. The recoveries obtained by this procedure were 97% or higher and were particularly good for nitrosamine levels of about 10 parts per 109. Tompkins and Griest [169, 170] have described a gas chromatographic method for the determination of N-nitrosodimethylamine in contaminated ground waters. This procedure utilizes a new solid-phase extraction procedure which extracts N-nitrodimethylamine at the nanogram per litre level from aqueous samples using a C (reversed18 phase) membrane extraction disc layered over a carbon-based extraction disc. The reversed-phase disc removes non-polar water insoluble neutrals and is set aside; the carbon-based disc is extracted with a small volume of dichloromethane. N-nitrosodimethylamine is quantified in the organic extract using a gas chromatograph equipped with both a short-path thermal desorber and a chemiluminescence nitrogen detector. The detection limit for the procedure is 3ng L-1 N-nitrosodimethylamine with a recovery of about 57%. A related procedure substitutes a standard automatic sampler for the short-path thermal absorber and is suitable for determining Nnitrosodimethylamine in heavily contaminated (greater than 300µg L-1) aqueous samples. The detection limit of this procedure is 300ng L-1 Nnitrosdimethylamine with a recovery of about 64%. 6.9.1.2 Nuclear magnetic resonance spectroscopy Fulton et al. [68] determined down to 510µg L-1 of N-nitrosodimethylamine in natural waters using 500MHz proton NMR spectroscopy. 6.9.2 Potable waters 6.9.2.1 Gas chromatography Fine et al. [69–71] have described a gas chromatographic method for the determination of N-nitroso compounds in potable water. A gas chromatograph equipped with a flame ionization detector and a 3.6m × 3mm stainless steel column packed with 10% diethyleneglycol succinate on 80–100 mesh Chromosorb W (HO) was used. Recoveries
Nitrosamines
449
obtained by this procedure were 77% or higher and were particularly good for the nitrosamine levels of about 10µg L -1 . Two different concentration and extraction procedures were used by these workers, one based on liquid–liquid extraction and the other based on the adsorption of the organic fraction on carbon and its subsequent extraction with chloroform and alcohol. In both cases, final quantitative analysis and identification were carried out on a single-column gas chromatograph equipped with N-nitroso compound specific thermal energy analyser. Figure 6.1 (a) is a chromatograph of 10µg of the mixture containing six N-nitroso compounds that were added to the water. Figure 6.1 (b) is the chromatogram of 10µL of the final dichloromethane extract following concentration on the Kuderna–Danish evaporator. At approximately 0.200µg L-1 concentration level the recovery efficiency is 30% for the dimethylnitrosamine, increasing to about 90% for the less volatile species. Detection limits ranged from 0.001µg L-1 (solvent extraction) to 10µg L-1 (carbon adsorption). Richardson et al. [72] have applied gas chromatography with a chemiluminescence detection system to the determination of microgram levels of nitrosamines (N-nitrosodimethylamine, N-nitrosodiethylamine, N-nitrosomorpholine and N-nitrosodiethanolamine, N-nitrosopyrrolidine, N-nitrosopiperidine and N-nitroso-5-methyl-1,3-oxazolidine) in potable water supplies. Nitrosamines may be removed from aqueous media by solvent extraction and subsequently concentrated by evaporation of the solvent, in order to detect levels as low as 0.01µg L-1. For the estimation of volatile dialkyl nitrosamines and Nnitrosopiperidine, N-nitrosopyrrolidine and N-nitrosomorpholine, 10N sulphuric acid was added to the sample which was then extracted with redistilled dichloromethane. 1.5M sodium hydroxide was added to the combined extract. After separation, the organic layer was dried over sodium sulphate and evaporated to 2.5mL at 46°C on a water bath. Hexane was added, and evaporation continued to about 250µL. Aliquots of 5µL were analysed for volatile nitrosamines using gas chromatography. In the chemiluminescence procedure, which detects all nitrosamines amenable to gas chromatography, effluent from a chromatograph passes into a catalytic chamber, whereupon the nitrosamine is fragmented to give rise to nitric oxide. This reacts with ozone and results in a chemiluminescent emission in the near infrared, which is detected with a photomultiplier tube. Interferences are minimized by placing a cold trap between the catalyst and ozone chamber, and by incorporating an optical filter in front of the photomultiplier. The presence and amounts of nitrosamines were confirmed in some of the samples using the gas chromatograph coupled to an AEI MS902
450
Nitrogen-containing compounds
Fig. 6.1 Gas chromatography of N-nitroso compounds in water, dimethylnitrosamine (DMN), diethylnitrosamine (DEN), dipropylnitrosamine (DPN), N-nitrosopiperidine (PIP), Nnitrosopyrrolidine (PYRN) and N-nitrosarcosinate (SARCOSN). (a) g.c. TEA spiking solution, (b) recovery from water at 0.010 L-1 level. (Reprinted with permission from Fine et al. [69]. Copyright (1975). Springer Verlag, New York.)
Nitrosamines
451
mass spectrometer. The nitrosamines were detected by parent-ion monitoring using peak matching, in the manner described by Gough and Webb [73]. The detection limit was 0.1µg L -1 for each of the nitrosamines. While measurement of the nitric oxide fragment by mass spectrometry is applicable to all nitrosamines, it results in a significantly poorer detection limit. Further, the mass spectrometer will respond to any compound giving rise to NO + , including C- and N-nitroso compounds, nitro compounds and nitramines. The upper limits of N-nitrosodimethylamine, N-nitrosodiethylamine, N-nitrosomorpholine and N-nitrosodiethanolamine detected were 0.2, 2.0 100 and 60µg L-1 respectively. 6.9.3 Waste waters 6.9.3.1 Gas chromatography–mass spectrometry Volatile nitrosamines have been determined in nanogram per litre amounts in waste water [74] by combined capillary gas chromatography– mass spectrometry. Dimethylnitrosamines, diethylnitrosamine and dipropylnitrosamine were determined. 6.9.4 Sewage effluents 6.9.4.1 Gas chromatography The gas chromatographic procedure described in Section 6.9.1.1 [67] for the determination of nitrosamines in natural waters has also been applied to sewage effluents. The gas chromatographic chemifluorescent detection procedure described by Richardson et al. [72] (Section 6.9.2.1) has been applied to the determination of microgram levels of nitrosamine (Nnitrosodimethylamine, N-nitrosodiethylamine, N-nitrosomorpholine, Nnitrosodiethanolamine, N-nitrosopyrrolidine, N-nitrosopiperidine, and N-nitroso-5-methyl-1,3-oxazolidine) imputs to sewage plants. 6.9.5 Trade effluents 6.9.5.1 Gas chromatography The Richardson et al. [72] gas chromatographic procedure (Section 6.9.2.1) has been applied to the determination of nitrosamines in trade effluents.
452
Nitrogen-containing compounds
6.9.6 High-purity waters 6.9.6.1 Gas chromatography Nitrosamines have been reported in deionized water by Fiddler et al. [75], Cohen and Backman [76] and Gough et al. [77]. N-nitrosodimethylamine and N-nitrosodiethylamine have been confirmed in water exposed to deionizing resins by Fiddler et al. [75]. Levels of N-nitrosodimethylamine detected were 0.03–0.34µg L-1 and N-nitrosodiethylamine 0.33– 0.83µg L-1. The highest levels of nitrosamines were found after resin regeneration. Nnitrosodimethylamine levels of 0.25µg L-1 and lower concentrations were also detected in deionized water by Cohen and Backman [76]. The origin of the nitrosamines in these reports was not identified. However, Gough et al. [77] suggested that N-nitrosodimethylamine present in the ionexchange resins (up to 125µg kg-1) as manufacturing contaminants, not the reaction of the components in the water with the resins, was responsible for the occurrence of trace levels of N-nitrodimethylamine (0.01µg L-1) detected in deionized water. N-nitrosodimethylamine was detected in the effluent when a 1M sodium nitrate solution was passed through a mixed strong anion and cation resin column by Angeles et al. [78]. Formation of N-nitrosodimethylamine was postulated by the cation acid-catalysed nitrosation of the amine–ammonium functional group on the strong anion resin during the deionization process. Kimoto et al. [5] attempted to ascertain whether one of the explanations offered by these earlier workers [77, 78] for the occurrence of nitrosamines in water deionized with strong cation and anion resins is correct or whether other factors are involved. They showed that Nnitrosodimethylamine was formed when tap water was passed through a column containing only the anion-exchange resin, thereby indicating that nitrosamine formation by the conventional acid-catalysed nitrosation reaction, as would be expected in the case of a mixed strong anion and cation resin system, was not the primary mechanism. The quaternary ammonium ion of the strong anion resin may be the amine precursor for nitrosamine formation. Strong anion and strong cation resins (Amberlite and Dowex brands) were ineffective in concentrating low levels of N-nitrosodimethylamine that were added to the influent. Accumulation of extremely low levels of nitrosamine already present in the water, therefore, also did not contribute importantly to the concentration of nitrosamine found in deionized water. In addition, the normal levels of cations and anions in water, and heavy metal ions were also not primarily responsible for this occurrence. There was, however, an unknown substance(s) in the tap water that promoted nitrosamine formation. This unknown soluble substance(s) can be removed by activated carbon treatment or degassing techniques.
Amino acids
453
Kimoto et al. [5] extracted nitrosamines from 1L water samples with 3 × 150mL portions of dichromomethane with shaking for 5min per extraction. The extracts were combined, dried over anhydrous sodium sulphate and concentrated in a Kuderna–Danish evaporator to 1.0mL. These workers determined nitrosamines in dichloromethane extracts by a gas chromatographic procedure. 6.9.6.2 Preconcentration Absorption on Ambersorb XE-34 cation-exchange resin and active carbon [79] has been used to preconcentrate nitrosamines from natural water prior to gas chromatography. 6.10 Amino acids 6.10.1 Natural waters 6.10.1.1 Gas chromatography Gardner and Lee [80] described an early gas chromatographic method for the estimation of dissolved free and dissolved total free and combined amino acids in lake water. The amino acids were first concentrated by ionand ligand-exchange chromatography. Volatile N-trifluoroacetyl methyl esters were prepared and determined on glass columns (4m × 3mm o.d.) packed with 0.7% of XE-60, 0.5% of OV-101, and 0.2% of QF-1 on Diatoport S, with flame ionization detection. Internal standards are added before derivatization. Recoveries were from 36 to 97% for the free acids and 55 to 93% for the hydrolysed samples. Recoveries were poor for phenylalanine and lysine. 6.10.1.2 Fluorimetry Gardner [81] isolated free amino acids at the 20nmol L-1 level in from as little as 5mL of sample, by cation-exchange, and measured concentrations on a sensitive amino acid analyser equipped with a fluorimetric detector. 6.10.2 Sea and estuary waters 6.10.2.1 High-performance liquid chromatography The classical work of Dawson and Pritchard [82] on the determination of a-amino acids uses a standard amino acid analyser modified to
454
Nitrogen-containing compounds
incorporate a fluorimetric detection system. In this method the samples are desalinated on cation-exchange resins and concentrated prior to analysis. The output of the fluorimeter is fed through a potential divider and low-pass filter to a compensation recorder. Dawson and Pritchard [82] point out that all procedures used for concentrating organic components from estuary or sea water, however mild and uncontaminating, are open to criticism, simply because of the ignorance as to the nature of these components in the sample. It is for instance feasible that during the process of desalting on ion-exchange resins under weakly acidic conditions, labile peptide linkages are disrupted or metal chelates dissociated and thereby larger quantities of ‘free’ components are released and analysed. An example of a chromatogram obtained from a saline sample and the mole percentage of each amino acid in the sample is depicted in Fig. 6.2. Mopper [83] has discussed developments in the reversed phase highperformance liquid chromatographic determination of amino acids in estuarine and sea water. He describes the development of a simple, highly sensitive procedure based on the conversion of dissolved free amino acids to highly fluorescent, moderately hydrophobic isoindoles
Fig. 6.2 Chromatogram of a saline extract (20mL) sample for amino acids collected at 6m in the Kiel Fjord. The concentrations of the individual acids were quantified as follows (nmol L-1): meto, 11; asp, 34.4; thr, 23.2; ser, 88; glu, 36; gly, 100; ala, 56; val, 16; ileu, 9.6; leu, 12; galactosamine and aminosugars, 4; tyr, 6.8; phe, 7.2; ß-ala, 20.8; ?-aminob. a, 14.4; orn, 44; lys, 12; hist, 7.2; arg. 8.6; cysSO H, 4; cit, trace; tau, cys, trace; glucosamine, trace; met, trace; urea, trace; 2 phosphoser, trace; OH-lys, trace. The total concentration of amino acid in the sample lies around 51 g L-1 assuming a mean molecular weight of 100. (Reprinted with permission from Dawson and Pritchard [82]. Copyright (1978). Elsevier Science Publishers, Netherlands.)
Chloroamino acids
455
by a derivatization reaction with excess o-phthaldehyde and a thiol, directly in sea water. Reacted samples were injected without further treatment into a reversed-phase high-performance liquid chromatography column, followed by a gradient elution. The eluted amino acid derivatives were detected fluorometrically. Detection limit for most amino acids was 0.1–0.2nmol per 500µL injection. Problems of inadequacies with the method itself, sample handling and whether chromatographically determined concentrations might be considered as biologically available concentrations in sea water, are discussed. 6.10.2.2 Fluorimetry Fluorimetry has been used to determine individual amino acids in marine waters [95, 96]. 6.10.2.3 Preconcentration Various other analytical techniques have been used for the determination of amino acids in sea water (Table 6.1). Separation on an automatic amino acid analyser working on the ion-exchange principle coupled with a flame fluorimetric detection has emerged as the method of preference. Bajor and Bohling [110] have investigated the use of piperidine for desalting amino acid extracts of sea water. 6.11 Chloroamino acids 6.11.1 Waste waters 6.11.1.1 Gas chromatography–mass spectrometry Chlorinated tyrosine has been determined [111] in municipal waste water by combined gas chromatography–mass spectrometry. 6.12 Nucleic acids 6.12.1 Natural waters 6.12.1.1 Spectrophotometric method Hicks and Riley [112] have presented a method for determining the natural levels of nucleic acids in lake waters, which involves preconcentration by adsorption on to hydroxyapatite, elution of the
Table 6.1 Amino acids in sea water
Notes FFA = free amino acids: CAA = combined amino acids.
458
Nitrogen-containing compounds
nucleic acids and then photometric determination of the ribose obtained from them by hydrolysis. 6.12.2 Sea water 6.12.2.1 Spectrophotometric method The spectrophotometric method [112] discussed above in Section 6.12.1.1 has been applied to sea water. 6.12.2.2 Titration method Seibert and Zahn [113] used precipitation titration with the cationic detergent N-cetyl-N-N,N-trimethylammonium bromide to quantify the nucleic acid content in activated sludge. With the introduction of toxins into the influent of a sewage treatment plant the ribonucleic acid content of the activated sludge fell sharply, and it was possible to correlate the deterioration in effective purification with a reduction in the ribo-nucleic acid fraction precipitate by N-cetyl-N,N,Ntrimethylammonium bromide. Advantages of the method are simplicity and speed. DNA and RNA can be determined in one operation and the process is not affected by the presence of dyes or compounds absorbing ultraviolet light. 6.13 Ethylenediamine tetraacetic acid (EDTA) EDTA salts are presented in low concentrations in detergent preparations and some food products. They are not biodegradable and might have an affect in mobilizing trace metals in river waters, i.e. in reducing their tendency to be removed from solution by adsorption and precipitation reactions and possibly causing desorption from contaminated river sediments. Hence, there is an interest in determining EDTA and its salts in river water, sewage and sewage effluents. 6.13.1 Natural waters 6.13.1.1 Gas chromatography Lockhart and Blakely [114] used gas chromatography to estimate ferric EDTA and its degradation products. Gas chromatographic analysis of the N-trifluoroacetyl-n-butyl esters of Fe(III)–EDTA and photolysis products was performed on a 4ft × 1/8 in i.d. (1.2m × 3mm) glass column of 3% QF-1 on 100/120 mesh Gas Chrom-Q using a Hewlett Packard Model
Ethylenediamine tetraacetic acid (EDTA)
459
7610A gas chromatograph with flame ionization detectors, with nitrogen as the carrier gas. Oven temperature was programmed from 110 to 260°C at 10° min-1, starting at 2min post-injection and holding for 2min at 260°C. Injection temperature was 250°C and detector temperature was 300°C. The N-trifluoroacetyl-n-butyl esters on the gas chromatographic column were identified by fragmentation pattern analysis on a Varian 1800 gas chromatograph coupled via a molecular separator to an AEIMS-30 double-beam mass spectrometer. Gardiner [115, 116] described a gas chromatographic method for the determination of EDTA in aqueous environmental samples. The separation of the major peaks is increased by preparing the ethyl derivatives of the sample compounds, 1,6-hexanediamine tetraacetic acid (HDTA) being used as internal standard. The lower limit of detection of the method is approximately 15µg L-1 with 25mL samples. The limit can be improved, if necessary, by using a larger sample volume. In this method the ethyl derivatives of the sample components were prepared so that the major peaks would be well separated. The ethyl esters of fatty acids up to and including the C 18 fatty acids eluted well before the EDTA derivatives and did not interfere. Fatty acids of greater chain length are invariably present at insignificantly low concentrations in environmental samples. It was also found that the linear analogue of EDTA, HDTA, when used as internal standard, did not interact with the humic acid present in the sample. A series of small peaks occurring in the region of the peak for HDTA did not interfere; these peaks may have originated from detergent compounds. Kari [171] and Schürch and Duberdorfer [172] both point out that gas chromatographic procedures need lengthy sample pretreatments. 6.13.1.2 Column chromatography Various workers [117, 173–175] have pointed out that only highperformance liquid chromatographic methods have sufficient sensitivity for the determination of EDTA. Nowack et al. [176] have described a procedure for the determination of dissolved EDTA in rivers and waste water treatment plants. The procedure is based on high-performance liquid chromatography. A distinction can be made between Fe(III)–EDTA and all other species. Nickel–EDTA can be detected semi-quantitatively. The fraction of EDTA adsorbed on to suspended particles or to sediment can be determined after desorption with phosphate. After complexation with Fe(III)– EDTA, the EDTA is detected by reversed-phase ion-pair liquid chromatography as the Fe(III)–EDTA complex at a wavelength of 258nm. The behaviour of a number of metal-EDTA complexes during
460
Nitrogen-containing compounds
analysis was checked. Fe(III)–EDTA was found to be the main species (60–70%); Ni–EDTA was less than 10% in most samples. 6.13.1.3 Polarography Dietz [118] used polarography to determine nitriloacetic acid, EDTA and other complexing agents in surface and ground waters using bismuth complexes at pH2. Concentrations of nitriloacetic acid and EDTA in the range 0.1–3mg L -1 could be determined selectively using cathode ray, impulse, or modified alternating-voltage polarography. Fayyad and Tatunji [119] described an indirect potentiometric stripping analysis method for determining EDTA as its 1:1 bismuth complex in natural waters. 6.13.1.4 Atomic absorption spectrometry Kunkel and Manahan [120] have described an atomic absorption method for determining strong heavy-metal chelating agents, such as EDTA and nitriloacetic acid in natural and waste waters. The method involved solubilization of cupric ions (added as 0.05M CuSO ) by the 4 chelating agents at pH10 in the boiling solution, filtration of the cool mixture, and then determination of cupric ions in the filtrate. The concentration of total strong heavy-metal chelating agents is proportional to the amount (in mg) of copper chelated in a standard volume of sample. 6.13.2 Sea water
6.13.2.1 Polarography Differential pulse polarography has also been used to determine EDTA and nitriloacetic acid in synthetic sea water and phytoplankton media [121]. Cadmium is used to convert a large fraction of either ligand to the reducible cadmium complex. The presence of competing metal cations, including copper, is not detrimental if the method of standard additions is used. The method was used in correlating the concentration of complexed and uncomplexed species of copper with phytoplankton productivity and the production of extracellular metalbinding organic compounds.
Ethylenediamine tetraacetic acid (EDTA)
461
6.13.3 Potable waters 6.13.3.1 Gas chromatography The gas chromatographic method described in Section 6.13.1.1 [114– 116] has been applied to the determination of EDTA in potable water. 6.13.4 Waste waters 6.13.4.1 High-performance liquid chromatography Dai and Helz [122] have described a liquid chromatographic method for the determination of down to 0.15µg L -1 of EDTA and related aminopolycarboxylic acids in waste waters. 6.13.5 Trade effluents 6.13.5.1 Liquid chromatography Dai and Helz [122] described a liquid chromatographic method for determining nitriloacetic acid, EDTA and related aminopolycarboxylic acids. An amperometric detector was used. 6.13.6 Sewage effluents 6.13.6.1 Gas chromatography Rudling [123] and Chau and Fox [124] have described a method in which the methyl ester was determined by gas–liquid chromatography using 1,2cyclohexanediamine tetraacetic acid as internal standard. This presented problems as the retention times of the methyl esters of EDTA and those of certain C fatty acids (stearic and oleic) which were also usually present in the 18 samples being investigated, were very similar and the peaks were not resolved under the conditions employed. It was considered unlikely that any stationary phase would completely resolve EDTA from the range of C fatty acids that 18 might be found in samples of sewage effluent. It was also found that humic acid in the sample, most of which failed to react with the esterifying agent, produced a greater than theoretical recovery of EDTA, apparently because of interaction between EDTA and non-reactive fraction of humic acid. The gas chromatographic procedure discussed in Section 6.13.1.1 [114–116] has been applied to the determination of EDTA in sewage effluents. Recoveries in spiked samples were between 79 and 136%.
462
Nitrogen-containing compounds
6.13.6.2 Miscellaneous Korsunovskii [125] has developed a procedure for determining very small amounts of EDTA, which can be used to analyse turbid and coloured solutions. The method is based on the proportionality of the photoelectric current generated in the photogalvanic cell to free EDTA concentration. 6.14 Nitriloacetic acid 6.14.1 Natural waters 6.14.1.1 Spectrophotometric method Coombs et al. [126] analysed mixtures of aminopolycarboxylic acids at the ppb level (US) by chemical kinetics. The method is based on the reaction of the nickel complexes of these acids with cyanide ion and the large differences in reaction rates for the formation of . The reaction is monitored spectrophotometrically (at 267, 285 or 310nm) with use of a stopped-flow system, and the results are calculated from a computer program that provides on-line data acquisition and performs a regressive differential kinetic analysis for the two components. The acids can be determined singly or up to three in admixture. The error is within ±5 to 10% for mixtures of ligands at the micromolar level and the sensitivity is 0.04µM (8 parts per 109) in water. 6.14.1.2 Gas chromatography Chau and Fox [124] concentrated nitriloacetic acid in lake water samples by passing them down a Dowex 1 column (formate form) and elution with 2.5M to 8M formic acid. The nitriloacetic acid is then esterfied, with heptadecanoic acid added as internal standard, by heating for 1h at 100°C in a sealed ampoule with propanol saturated with hydrogen chloride. The propyl esters are analysed on a stainless steel column (6ft × 0.25in (1.8m × 6mm)) packed with 3% of OV-1 on Chromosorb WHP (80–100 mesh), temperature programmed from 180 to 225°C min-1, and operated with nitrogen as carrier gas (65mL min-1) and flame ionization detection. The calibration graph is rectilinear for up to 20µg of nitriloacetic acid as ester. The limit of detection is 0.01µg and at the level of 20µg L-1 the standard deviation was 1.33µg L-1 and the coefficient of variation was ±6.3%. Murray et al. [127] converted nitriloacetic acid to its trimethyl ester prior to gas chromatographic determination on a column packed with
Nitriloacetic acid
463
2% of poly(ethanediol) on Chromosorb W. Down to 25µg L-1 nitriloacetic acid could be determined by this procedure. Warren and Malec [128] determined nitriloacetic acid and related aminopolycarboxylic acids (iminodiacetic acid, glycine and sarcosine) in inland waters and sewage effluents by converting to the butyl or the Ntrifluoracetyl esters followed by chromatography on dual glass Ushaped columns (1.9m × 2mm) packed with 0.65% of ethanediol adipate on acid-washed Chromosorb W (80–100 mesh), temperatureprogrammed from 80 to 220°C and operated in the differential mode with flame ionization detectors. The signal was fed to a digital integrator and then to both channels of a dual-pen recorder. 6.14.1.3 Polarography Wernet and Wahl [129] removed interfering cations and heavy metals that form complexes with nitriloacetic acid from surface water and effluent samples by equilibrating at pH3 with Dowex 50W-X8 resin (sodium form) prior to polarography in the presence of ammoniacal cadmium buffer solution at pH8. Haberman [130] determined down to 0.02ppm of nitriloacetic acid in water and sewage by first passing the sample, adjusted to pH3, through a cation exchanger. After adjustment to pH7.0, the percolate is then passed through an anion exchanger to absorb the nitrilotriacetate, which is subsequently eluted with sodium chloride in 0.1M acetic acid buffer of pH4.7. Nitriloacetic acid is then determined by adding a known amount (in excess) of trivalent indium to the solution and measuring the height of the wave due to reduction of the In 3+nitriloacetic acid complex at -0.79V vs the SCE. An isotope dilution technique, usually 14C-labelled nitriloacetic acid, is used to correct for incomplete recovery. Low concentrations (1–10ppm) of nitriloacetic acid have been determined in lake water by a method [131] which involved conversion to its 1:1 cadmium complex. The resulting solution (adjusted to pH9) is subjected to polarography between -0.25 and 1.2V (vs the SCE) in 0.1M potassium nitrate, as supporting electrolyte; Cd2+ gives a wave at 0.060V and the Cd–nitriloacetic acid complex gives a wave at 0.97V. To determine nitriloacetic acid in submicromolar concentrations it is advantageous to use a pH of 8.5 to avoid precipitation of cadmium hydroxide. Afghan et al. [132] developed an automated method for the determination of nitriloacetic acid in natural water and sewage samples. The method is based on the formation of the bismuth nitrilotriacetic acid complex at pH2 followed by determination by twin cell oscillographic DC polarography. As little as 10µg L-1 of
464
Nitrogen-containing compounds
nitrilotriacetic acid can be determined, with no preconcentration of the sample being required. The coefficient of variation for 100mg L-1 was 1.3%. Dietz [118] used polarography to determine nitriloacetic acid, EDTA and other complexing agents in surface and ground waters using bismuth complexes at pH2. Concentrations of nitriloacetic acid and EDTA in the range 0.1–3mg L-1 could be determined selectively using cathode ray, impulse, or modified alternating voltage polarography. Haring and Van Delft [133] studied the application of derivative pulse polarography at a hanging mercury drop electrode to the determination of nitriloacetic acid in water. Voulgarpoulos et al. [134] carried out indirect determinations of nitriloacetic acid and EDTA in natural waters by differential pulse anode stripping voltammetry. Both substances are determined by complex formation with excess trivalent bismuth ions at pH2 followed by back-titration of the excess using a hanging mercury drop electrode at a potential of -0.15V. The peak current due to uncomplexed bismuth is recorded. By means of a second deposition at -0.35V, the bismuth from the Bi-nitriloacetic complex can be recorded independently of that accounted for by EDTA. The lower limit of determination is 0.2µg L-1 nitriloacetic acid for the deposition time of 2min. Interference due to ferric ions may be eliminated by the addition of ascorbic acid as a reducing agent, but copper ions, if present in excess of 40µg L -1 , must first be removed by preelectrolysis with a mercury pool electrode. 6.14.1.4 Potential sweep potentiometry Afghan and Goulden [135] used linear potential sweep chronoamperometry of the nitriloacetic acid–lead complex to determine down to 10µg L-1 of nitriloacetic acid in water. To determine total nitriloacetic acid, the sample is acidified to pH1 to release the nitriloacetic acid from heavy metal complexes and these metals are masked by adding EDTA before bringing the pH back to 8 to form the lead complex. Large amounts of bis-(2-aminoethoxyl)-ethaneN,N,N’,N’-tetraacetic acid interfere with the lead–nitriloacetic acid method. 6.14.1.5 Atomic absorption spectrometry The atomic absorption spectrophotometric method [120] described in Section 6.13.1.4 has been applied to the determination of nitriloacetic acid in natural waters.
Nitriloacetic acid
465
6.14.2 Sea water 6.14.2.1 Polarography The differential pulse polarographic method [121] discussed in Section 6.13.2.1 has been applied to the determination of nitriloacetic acid in sea water. 6.14.3 Potable waters 6.14.3.1 Gas chromatography Aue et al. [136] determined parts per billion (US) of nitriloacetic acid and citric acid in tap water and sewage effluents. Following a preliminary clean-up and concentration procedure the acids are converted to their butyl esters and chromatographed on a Carbowax on Celite 545 column at 183°C. By converting nitriloacetic acid to its trimethylsilyl ester, Stolzberg and Hume [137] were able to extend the limit of detection of this substance down to 1µg L-1 without a concentration step. Williams et al. [138] applied a nitrogen-specific detector to a survey of the levels of nitriloacetic acid to its tri-n-butyl ester in US tap water supplies at concentrations approaching 1ppb (US). These workers based their method on that described by Aue et al. [136], described earlier. Williams et al. [138] used a Perkin–Elmer Model 910 gas chromatograph, equipped with a single column, a two-way effluent splitter, a flame ionization detector and a nitrogen–phosphorus detector operating in the nitrogen mode. The column was 6ft × 1/4 in. o.d. (1.8m × 6mm) glass, packed with either 5% OV-101 or 3% OV-210 on 80–100 mesh Chromosorb WHP. The carrier gas was helium at a flow rate of 60mL min-1 and the effluent splitter diverted 60% to the flame ionization detector and a 40% to the nitrogen detector. Hydrogen and air flows were optimized for each detector. The injector and detector temperatures were 240 to 280°C respectively and the column and interface temperatures 200 and 250°C. Aue et al. [136] claimed a limit of detection of 1ppb nitriloacetic acid for a 50mL water sample, but preliminary investigations of Williams et al. [138] with standard solutions of the tri-n-butyl ester showed that quantitation at this level was difficult due to interference from the solvent peak when using acetone as the injection solvent as specified by Aue et al. [136] The use of alternative injection solvents gave some improvement but quantitation was still difficult. Analysis of standard solutions of the tri-n-butyl ester, equivalent to 1ppb nitriloacetic acid in
466
Nitrogen-containing compounds
a 50mL water sample, showed that the sensitivity of the nitrogenspecific detector was adequate, quantitation was straightforward, and there was minimal interference from the injection solvent, acetone. The nitrogen-selective detector gave a linear response over the range of 1– 1000ng injected of the tri-n-butyl ester of niriloacetic acid. The isolation procedure of Aue et al. [136] gave a satisfactory chromatogram for a control blank water sample provided that all solvents were redistilled in glass and the ion-exchange resin and glassware were thoroughly washed before use. The lower detection limit is four times the level of the blank which would give a detection limit of ca 0.2ppb nitriloacetic acid for a 500mL water sample. Recoveries of nitriloacetic acid from water samples spiked with 1–1000ppb nitriloacetic acid were greater than 90%. Reichert and Linckens [139] have reviewed gas chromatographic methods for the determination of nitriloacetic acid in potable waters. They point out that esterification of the nitriloacetic acid is required to enable it to be volatilized in the gas–liquid chromatographic column, and they compared a range of esterification reagents and conditions for simplicity and speed of operation. The method chosen involved treatment of a concentrated sample with a mixture of n-propanol– acetyl chloride (10:1) and the resulting nitroacetic acid–propyl ester injected into a column, which was fitted with a nitrogen-sensitive detector. The detection limit for nitriloacetic acid in potable water is about 1µg L-1. 6.14.3.2 Fluorimetry Robinson and Lott [140] used a fluorimetric method for the determination of down to 0.2ppb nitriloacetic acid in tap water. The method is based on the displacement of 8-hydroxyquinoline from its fluorescent complex with Ga3+ by nitriloacetic acid, the gallium complex of which is not fluorescent. The reaction is carried out in a medium buffered by acetate at pH6 and containing hydroxylammonium chloride (to reduce Fe3+) and KI–NaH PO –NaCN masking reagent. Standard 2 4 additions of nitriloacetic acid are made, and the organogallium complexes are extracted into chloroform for fluorescence measurement (and thereby separated from the 8-hydroxyquinoline complexes of magnesium and calcium). 6.14.3.3 Polarography Fayyad [141] described an indirect determination of nitriloacetic acid in potable waters using bismuth nitriloacetic acid complex with potentiometric stripping analysis. Spiked samples containing 6–255µg L-1
Nitriloacetic acid
467
of nitriloacetic acid were adjusted to pH2–2.3 (hydrochloric acid), a solution of known bismuth (III) concentration was added to give final concentration which exceeded that of nitriloacetic acid by 1.5–2.3 times, then 58µM of free bismuth (III) and 0.42µM of potassium dichromate (oxidant) were added. The electrode system (mercury on glass carbon) was immersed in the solution after stirring for 10min and voltage stripping curves recorded. Two subsequent time curves were obtained by adding 50µL of bismuth (III) (10mg L-1) solution. Each time nitriloacetic acid concentrations were calculated from the difference between concentrations of added bismuth (III) and that of free bismuth determined by potentiometric stripping analysis. 6.14.4 Waste water 6.14.4.1 Titration method Sekerka et al. [142] described a thallium nitrate potentiometric titration procedure for the determination of down to 0.1µg L-1 of nitriloacetic acid in waste water and sewage. To the sample (100mL) was added 0.01M thallium nitrate (0.1mL) and 10M sodium hydroxide, the mixture was filtered, the filtrate adjusted to pH7 by addition of 10M nitric acid and hexamine (2g) and 0.01M ammonium pyrrolidine-1-carbodithioate added (this preliminary treatment removes interfering cations and reducing substances). The solution was titrated with 0.1M thallium nitrate (Tl3+ forms a 1:2 complex with nitriloacetic and 1:1 complexes with EDTA) with the use of a combination platinum redox electrode (Orion 96-78). A second 100mL sample is treated similarly, but in the first step 0.1M cupric nitrate is added in place of the thallium nitrate and titrated with 0.1mM cupric nitrate (Cu2 + forms 1:1 complexes with all the complexing agents tested). The titration curves exhibit two end points corresponding to: 1 2
T13+ or (Cu2+) consumed by ammonium pyrrolidine-1-carbodithioate; Nitriloacetic acid (or other ligands) present in the sample.
The content of nitriloacetic acid is calculated from the differences in titres. The recovery of nitriloacetic acid admixture with EDTA in synthetic sewage samples ranges from 90 to 102%. 6.14.4.2 Liquid chromatography Dai and Helz [122] have described a liquid chromatographic procedure for the determination of down to 0.1mg L-1 of nitriloacetic acid in waste waters.
468
Nitrogen-containing compounds
6.14.4.3 Atomic absorption spectrometry The atomic absorption spectrophotometric method [120] described in Section 6.13.1.4 has been applied to the determination of nitriloacetic acid in waste waters. 6.14.4.4 Miscellaneous Kirk et al. [143] determined nitriloacetic acid in waste waters by gas chromatography, differential pulse polarography and spectrophotometry. Gas chromatography gave good results down to 10µg L-1. Differential pulse polarography was quicker and was more precise for all sample types, but only down to 100µg L-1 nitriloacetic acid could be determined. The detection limit of the spectrophotometric method was 500µg L-1. 6.14.5 Trade effluents 6.14.5.1 Liquid chromatography Dai and Helz [122] have described a liquid chromatographic procedure for the determination of nitriloacetic acid in trade effluents. 6.14.6 Sewage effluents 6.14.6.1 Titration method The potentiometric titration procedure [142] discussed in Section 6.14.4.1 has been applied to the determination of nitriloacetic acid in sewage effluents. 6.14.6.2 Gas chromatography Rudling [144] determined nitriloacetic acid gas chromatographically at the 0.1µg L-1 level in sewage. Following separation of the nitriloacetic acid on an anion-exchange column, it is derivatized with boron trifluoride in 2-chloroethanol to produce 2-chloroethylnitrilotriacetate and gas chromatographed on a column packed with 2% QF-1 on Varaport 30. Detection is achieved by electron capture. Chloride, sulphate, calcium and magnesium did not interfere in this procedure. Rudling [144] simultaneously determined 0.0–10.02µg of nitriloacetic acid, EDTA and diethylenetriamine pentaacetic acid in sewage samples as their methyl esters by gas chromatography. The sample, adjusted to pH7, is extracted with chloroform, 1,2-diaminocyclohexane tetraacetic
Nitriloacetic acid
469
acid, an internal standard is added and the mixture treated with methanolic boron trifluoride. Chloroform extracts are injected into a column packed with 5% V-17 on Aeropak, temperature programmed from 150 to 285°C at 10min-1 using helium as carrier gas. Cadmium, copper, zinc and nickel, and iron did not interfere. Aue et al. [136] determined micrograms per litre of nitriloacetic acid and citric acid in sewage effluents. Following a preliminary clean-up and concentration the acids are converted to their butyl esters and chromatographed on a Carbowax on Celite 545 column at 183°C. The gas chromatographic procedure [128] discussed in Section 6.14.1.2 has been applied to the determination of nitriloacetic acid in sewage effluents. 6.14.6.3 Ultraviolet spectrometry Longbottom [145] used high-speed ion-exchange chromatography to determine nitriloacetic acid with a sensitivity of 1µg L-1. Nitriloacetic acid was separated on a stainless steel column packed with SAX, a strong anion-exchange resin, coated on Zipax. The mobile phase was 0.02M Na B O (pH9). Possible interference from metal ions was overcome by 2 4 7 converting all metal–nitriloacetic acid complexes into Fe3+- nitriloacetic acid complex. The nitriloacetic acid was monitored at 254nm with an ultraviolet spectrophotometric detector; maintenance of the pH at 9 was essential because the extinction of nitriloacetic acid at this wavelength varies widely with pH. The method was applied to the analysis of sewage samples. 6.14.6.4 Polarography The polarographic method [130] discussed in Section 6.14.1.3 has been applied to the determination of nitriloacetic acid in sewage. The automated polarographic method [132] discussed in Section 6.14.1.3 has been applied to the determination of nitriloacetic acid in sewage. 6.14.6.5 Miscellaneous Longman et al. [146] compared five procedures for removing interferences in the determination of nitriloacetic acid in sewage effluents. None of these methods completely overcame interference by metals. These workers then developed an alternative procedure based on passing the filtered sample through a column of Chelex 100 chelating resin (Na + form). To the eluate was added zircon reagent and the
470
Nitrogen-containing compounds
Table 6.2 Miscellaneous nitrogen compounds
References
471
extinction was measured at 620nm. The method was applicable in the range 0–10µg L-1 nitriloacetic acid in sewage. 6.15 Other nitrogen compounds Methods for the determination of other types of nitrogen compounds, including heterocyclics, N-methylpyrrolidone, pyridazinones, 1-methyl2-pyrrolidine, guanazoles, aminotriazoles, alkylhydrantoins, diazo compounds, azarenes, N-chloramines, diethanolamine, monoalkyl quaternary compounds, hydrazine, chlorinated isocyanurates, ecaprolactam, ureas and dodecyl guanidenes are reviewed in Table 6.2. References 1 Florence, T.M. and Farner, Y.J. Anal. Chim. Acta 63 255 (1973) 2 Hermanson, H.P., Helrich, K. and Carey, W.F. Analytical Letters (London) 1 941 (1968) 3 Metera, J., Rabhl, V. and Mostecky. J. Water Research 10 137 (1976) 4 Onuska, F.I. Water Research 7 835 (1973) 5 Kimoto, W.I., Dooley, C.J., Canoe, J. and Fiddler, W. Water Research 14 869 (1980) 6 Avery, M.J. and Junk, G.A. Analytical Chemistry 57 790 (1985) 7 Dyat-Lovitskaya, F.G. and Botvinova, L.E. Khim. Volokna. 1 64 (1970). Ref. Z. Khim. 19GD (13) Abstr. No. 13G308 (1970) 8 Nishikawa, Y. Journal of Chromatography 349 349 (1987) 9 Yang, X.-H., Lee, C. and Scranton, M.I. Analytical Chemistry 65 857 (1991) 10 Fishman, G.I. and Pevzner, J.D. Zavod. Lab. 36 926 (1970) 11 Dyat-Lovitskaya, F.G. and Botvinova, L.E. Khim. Volokna. 1 64 (1970). Ref. Z. Khim. 19GD (13) Abstr. No. 13G308 (1970) 12 Sherma, J., Chandler, K. and Ahringer, J. Journal of Liquid Chromatography 7 2743 (1984) 13 Sherma, J. and Pallasta, L. Journal of Liquid Chromatography 9 3439 (1986) 14 El-Dib, M.A., J. Assoc. Anal. Chem. 54 1383 (1971) 15 Wegman, R.C.C. and DeKorte, G.A.L. Int. J. Environ. Anal. Chem. 9 1 (1981) 16 Wanatabe, T., Hongu, A., Honda, K., Nakazato, M., Kanno, M. and Saitoh, S. Analytical Chemistry 56 251 (1984) 17 Stuber, H. and Leenheer J.A. Analytical Chemistry 55 111 (1983) 18 Nielen, M.W.F., Frei, R.W. and Brinkman, U.A.T. Journal of Chromatography 317 557 (1984) 19 Varney, M.S. and Preston, M.R. Journal of Chromatography 348 265 (1980) 20 Zatkovetskii, V.M., Safonova, Z.B., Koren’kov, V.N., Nevskii, A.B. and Sosin, S.L. Zavod. Lab. 37 1434 (1971) 21 Bark, L.S., Cooper, R.L. and Wheatstone, K.C. Water Research 6 117 (1972) 22 Jenkins, R.L. Bulletin of Environmental Contamination and Toxicology 13 436 (1975) 23 American Public Health Association. Supplement to the 15th Edition of Standard Methods for the Examination of Water and Waste Water. Method 111, p. 548(1975) 24 Riggin, R.M., Cole, T.F. and Billets, C. Analytical Chemistry 55 1862 (1983) 25 Di Corcia, A. and Samperi, R. Analytical Chemistry 62 1490 (1990)
472
Nitrogen-containing compounds
26 Polishchuk, O.M. and Gorbonds, T.V. Soviet J. Chem. Technol. 5 50 (1983) 27 Coulte, L.T., Hargescheinger, E.E., Pascuto, F.M. and Baker, G.B. Journal of Chromatographic Science 19 151 (1981) 28 Schulz, B. J. Chromat 299 484 (1984) 29 Heller, C.A., Greni, G.R. and Erikson, E.D. Analytical Chemistry 54 286 (1982) 30 Leggett, D.C. Analytical Chemistry 49 880 (1977) 31 Fan, T.Y., Ross, R., Fine, D.H. and Keith, L.H. Environmental Science and Technology 12 692 (1978) 32 Patil, S.S. Analytical Letters (London) 21 1397 (1988) 33 Hess, T.L., Gurdry, L.J. and Sibley, S.D. Bull. Environ. Contam. Toxicol. 13 579 (1975) 34 Walsh, J.T., Chalk, R.C. and Merritt, C. Analytical Chemistry 45 1215 (1973) 35 Heller, C.A., McBride, R.R. and Ronning, M.A. Analytical Chemistry 49 2251 (1977) 36 Jenik, J., Renyer, F. and Smetanova, M. Vodni. Hospodarstvi. Series B. 31 325 (1981) 37 Richard, J.J. and Junk, G.A. Analytical Chemistry 58 723 (1986) 38 Belkin, F, Bishop, R.W. and Sheely, G.A. Journal of Chromatographic Science 23 532 (1985) 39 Spanggord, R.J., Gibson, B., Keck, R.G. and Thomas, D.W. Environmental Science and Technology 16 229 (1982) 40 Maskarinec, M.P., Manning, D.L., Harvey, R.W., Greist, W.H. and Tomkins, B.A. Journal of Chromatography 302 51 (1984) 41 Bauer, F., Grant, C.L. and Jenkins, F.F. Analytical Chemistry 58 176 (1986) 42 Spangoord, R.J. and Suta, B.F. Environ. Sci. Technol. 16 233 (1982) 43 Cranmer, M. Bulletin of Environmental Contamination and Toxicology 5 329 (1970) 44 Bengtsson, G. Journal of Chromatographic Science 23 397 (1985) 45 Chambou, P. and Chambon, R. Journal of Chromatography 87 287 (1973) 46 Hoffsammer, J.C., Glover, D.J. and Hazzard, C.V. Journal of Chromatography 195 435 (1980) 47 Makarova, C.U. Cig. Sanit. 5 61 (1971). Ref. Zh. Khim. 19GD. (20) Abstr. No. 20G191 (1971) 48 Stefanescu, T. and Ursu, G. Materiale Plate. 10 330 (1973) 49 Ghersin, Z., Stitzi, H. and Wanea, R. Revta. Chim. 20 689 (1969) 50 Warren, J.M. and Beasley, R.K. Analytical Chemistry 56 1963 (1984) 51 Arkell, G.H. and Croll, B.I., The Water Research Association, Medmenham, Harlow, UK, Report T P78. The Determination of Acrylamide in Water, December (1970) 52 Croll, B.T. and Simpkins, G.M. Analyst (London) 97 281 (1972) 53 Hashimoto, A. Analyst (London) 101 932 (1976) 54 Husser, E.R., Stehl, R.H., Price, D.R. and DeLap, R.A. Analytical Chemistry 49 154 (1977) 55 Ludwig, F.J. and Besand, M.F. Analytical Chemistry 50 185 (1978) 56 Brown, L. and Rhead, M. Analyst (London) 104 391 (1979) 57 Croll, B.T., Arkell, G.M. and Hodge, R.P. Water Research 8 989 (1974) 58 Ortubia, M.G. and Mateos, F.L. Technologia Del Agua 36 27 (1987) 59 Parazak, D.P., Burkhardt, C.N. and McCarthy, K.J. Analytical Chemistry 59 1444 (1987) 60 Standing Committee of Analysts, H.M. Stationery Office, London. Methods for the Examination of Waters and Associated Materials (1988). Determination of Acrylamide Monomer in Waters and Polymers (1987)
References
473
61 Ludi, H., Garm, D., Mchisalba, O., Chisalba, O. and Schar, H.P. International Journal of Environmental Analytical Chemistry 33 131 (1988) 62 Brown, L., Rhead, H.H. and Bancroft, K.G.C. Analyst (London) 107 749 (1982) 63 Leung, R.W.M., Pandey, R.N. and Das, B.S. Environmental Science and Technology 21 476 (1987) 64 Jungreis, E. Analytical Letters (London) 14 1177 (1971) 65 Bezazyan, I.I., Kulikova, O.I. and Wicheva, I.A. Zavod. Lab. 38 415 (1972) 66 Mills, A.I. and Alexander, M.J. Environmental Quality 5 437 (1976) 67 Nikaido, M.M., Raymond, D.D., Francis, A.J. and Alexander, M. Water Research 11 1085 (1977) 68 Fulton, D.B., Sayer, B.G., Bain, A.D. and Malle, H.V. Analytical Chemistry 64 349 (1992) 69 Fine, D.H., Rounbehler, D.P., Huffman, F. and Epstein, S.S. Bull. Environ. Contam. Toxicol. 14 404 (1975) 70 Fine, D.H., Lieb, D. and Rounbehler, D.P. Unpublished work 71 Fine. D.H. and Rounbehler, D.P. Unpublished work 72 Richardson, M.L., Webb, K.S. and Gough, T.A. Ecotoxicol. Environ. Safety 4 207 (1980) 73 Gough, T.A. and Webb, K.S. Journal of Chromatography 79 57 (1973) 74 Hartmetz, G. and Shemrova. J. Bull. Environmental Contamination and Toxicology 25 106 (1980) 75 Fiddler, W., Pensabene, J.W., Doerr, R.C. and Dooley, C.J. Food Cosmet. Toxicol. 15 441 (1977) 76 Cohen, J.B. and Backman, J.D. Environmental Aspects of N-Nitroso Compounds (edited by Walker, E.A. et al.), pp. 257–372, International Agency for Research on Cancer (1978) 77 Gough, T.A., Webb, K.S. and McPhail, M.F. Food Cosmet. Toxicol. 15 437 (1977) 78 Angeles, R.M., Keefer, L.K., Roller, P.P. and Uhm, S.J., Environmental Aspects of N-Nitroso Compounds (edited by Walker, E.A. et al.), pp. 357–372, International Agency for Research on Cancer (1978) 79 Kimoto, W, Dooley, C.J., Carre, J. and Fiddler, W. Water Research 15 1099 (1981) 80 Gardner, W.G. and Lee, G.F. Environmental Science and Technology 7 719 (1973) 81 Gardner, W.G. Marine Chemistry 6 15 (1978) 82 Dawson, R. and Pritchard, R.G. Marine Chemistry 6 27 (1978) 83 Mopper, K. Sci. Total Environment 49 115 (1986) 84 Tatsumoto, M., Williams, T.J., Prescott, J.M. and Hood, D.W. Journal of Marine Research 19 89 (1961) 85 Palmork, K.H. Acta Chem. Scand. 17 1456 (1963) 86 Degens, E.T., Reuter, J.H. and Shaw, N.F. Geochim. Cosma. Acta 28 45 (1964) 87 Rittenberg, S.C., Emery, K.O., Hulsemann, J., Degens, E.T., Fay, R.C., Reuter, J.H., Grady, J.R., Richardson, S.H. and Bray, E.E. Sediment. Petrol 33 140 (1963) 88 Siegel, A. and Degens, E.G. Science 151 1098 (1966) 89 Webb, K.L. and Wood, L., In: Scova, N.B. (ed.), Automation in Analytical Chemistry (Technicon Synposium 1966 ), Mediad Incorporated, New York, 1 440–444 (1967) 90 Hobbie, J.E., Crawford, C.C. and Webb, K.L. Science 159 1463 (1968) 91 Palmork, K.H. Int. Counc. Explor. Sea. Hydrography Committee, C16CM (1969)
474
Nitrogen-containing compounds
92 93 94 95 96 97 98 99 100 101 102
Starikova, N.D. and Korzhikova, R.I. Oceanol. 9 509 (1969) Bohling, H. Mar. Biol. 6 213 (1970) Riley, J.P. and Seagar, D.A. J. Mar. Biol. Assoc. U.K . 50 713 (1970) Andrews, P. and Williams, P.J.LeB. J. Mar. Biol. Assoc. U.K. 51 11 (1971) Pocklington, R., Nature (London) 230 374 (1971) Clark, M.E., Jackson, G.A. and North, W.J. Limnol Oceanogr. 17 749 (1972) Bohling, H. Marine Biology 16 281 (1970) Coughenower, D.D. and Curl, H.C. Jr. Limnol. Oceanogr. 20 128 (1975) North, B.B. Limnol. Oceanogr. 20 20 (1975) Crawford, C.C., Hobbie, T.E. and Webb, K.L. Ecology 55 551 (1974) Brockmann, U.H., Eberlein, K., Junge, H.D., Majer-Reimer, E., Siebers, D. and Trageser, H. Entwichlung naturlicher Planktonpopulationen in einem outdoor-Tank mit nahrstoffarmem Meerwasser . I I. Konzentrationsveranderungen von gelosten neutralen Kohlenhydraten und freien gelosten Aminosauren. Berichte aus dem Sonderforschungsbereich Meeresforschung. SFB94 . Universitat Hamburg, 6 166–184(1974) Williams, P.J.LeB., Berman, T. and Holm-Hansen, O. Marine Biology. 35 41 (1976) Garrasi, C. and Degens, E.T., Analytische Methoden zur saulenchromatographischen Bestimmung von Arminosauren und Zuckern im Meerwasser und Sediment. Berichte aus dem Projeekt DFG-DE74/ 3 : LitorolforschungAbwasser in Kustennahe, DEG-Abschlusskolloquium, Bremerhaven (1976) Dawson, R. and Mopper, K. Anal. Biochem. 83 100 (1977) Lee, C. and Bada, J.L. Earth Planet Sci. Lett. 26 61 (1975) Lee, C. and Bada, J.L. Limnol. Oceanog. 22 502 (1977) Daumas, R.A. Marine Chemistry 4 225 (1976) Dawson, R. and Gocke, K. Oceanol. Acta 1 80 (1977) Bajor, M. and Bohling, H.Z. Analytical Chemie 249 190 (1970) Burleson, J.L., Peyton, G.R. and Glaze, W.H. Environ. Sci. Technol. 14 1354 (1980) Hicks, E. and Riley, J.P. Anal. Chem. Acta, 116 137 (1980) Seibert, G. and Zahn, R.K. Gas u. Wasserfach. (Wasser) Abwasser 117 184 (1976) Lockhart, H.B. and Blakely, R.W. Environmental Science and Technology 9 1035 (1975) Gardiner, J. Analyst (London) 102 120 (1977) Gardiner, J. Water Research Centre, Stevenage Laboratory, Herts, UK, Technical Memorandum TM101 (1975) Venezby, D.L. and Rudzinski, W.E. Analytical Chemistry 56 315 (1984) Dietz, F.Z. Wasser Abwasser Forsch. 7 74 (1974) Fayyad, M. and Tatunji, Z. Analytical Letters (London) 21 1425 (1988) Kunkel, R. and Manahan, S.E. Analytical Chemistry 45 1465 (1973) Stolzberg, P.J. Anal Chim. Acta 92 139 (1977) Dai, J. and Helz, G.R. Analytical Chemistry 60 301 (1988) Rudling, L. Water Research 6 871 (1972) Chau, A. and Fox, M.E. Journal of Chromatographic Science 9 271 (1971) Korsunovskii, G.A. Z. Analytische Chemie 29 1244 (1974) Coombs, L.O., Vasiliades, J. and Margerum, P.W. Analytical Chemistry 44 2235 (1972) Murray, D., Povoledo, D. and Fish, J. J. Research Board, Canada 28 1043 (1971) Warren, C.B. and Malec, E. Journal of Chromatography 64 219 (1972) Wernet, J. and Wahl, K. Z. Analytische Chemie 251 373 (1970)
103 104
105 106 107 108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126 127 128 129
References
130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162 163 164 165 166 167
475
Haberman, J.P. Analytical Chemistry 43 63 (1971) Asplund, J. and Wanninen, E. Analytical Letters (London) 4 267 (1971) Afghan, B.K., Goulden, P.D. and Ryan, J.P. Analytical Chemistry 44 354 (1972) Haring, J.A. and Van Delft, W. Analytical Chemistry 94 201 (1977) Voulgarpoulos, A., Valenta, P. and Nurnberg, H.W. Fresenius Z. für Analytische Chemie 317 367 (1984) Afghan, B.K. and Goulden, P.D. Environmental Science and Technology 15 60 (1971) Aue, W.A., Hastings, C.R., Gerhardt, K.O., Pierce, J.D., Hill, H.M. and Moseman, R.F. Journal of Chromatography 72 259 (1972) Stolzberg, R.G. and Hume, D.N. Journal of Chromatography 49 374 (1977) Williams, P.J., Benoit, F, Muzeka, K. and O’Grady, R. Journal of Chromatography 136 423 (1977) Reichert, J.K. and Linckens, A.H.M. Environ. Technol. Lett. 1 42 (1980) Robinson, J.L. and Lott, P.F. Microchemical Journal 18 128 (1973) Fayyad, M. Analytical Chemistry 59 209 (1987) Sekerka, I.L., Lechner, J. and Afghan, B.K. Analytical Letters (London) 6 977 (1973) Kirk, A.W.W., Perry, R and Lester, J.N. International Journal of Environmental Analytical Chemistry 12 293 (1982) Rudling, L. Water Research 5 831 (1971) Longbottom, J.E., Analytical Chemistry 44 418 (1972) Longman, G.F., Stift, M.J. and Gardiner, D. K. Water Research 5 1171 (1971) Kupor, V.G., Nauch. Trudy Omsk. Med. Dist. 160 (88) (1968). Ref. Zh. Khim. 19GD Abstr. No. 24G203 (1969) Jaroslav, N. Chemiky. Prumsyl. 20 575 (1970) Scoggins, M.W. and Skurcenski, L.J. Chromatographic Science 15 573 (1977) Matisova, E., Lehotay, J., Garaj, J. and Violova, A. Journal of Chromatography 237 164 (1982) Fedonina, V.F., Fedyainov, N. V and Agapova, S.A. Soviet Journal of Water Chemistry Technology 3, 57 (1981) Stevens, R. Analytical Chemistry 56 1608 (1984) Alary, J., Barban, C., Escrient, C. and Vandacle, J. Environmental Technology Letters 4 93 (1985) Olson, E.S., Diehe, W. and Miller, D.J. Analytical Chemistry 55 1111 (1983) Bruins, A.P., Weidaft, L.O.G., Henion, J.D. and Budde, W.L. Analytical Chemistry 59 2647 (1987) Steinheimer, T.R. and Ondrus, M.G. Analytical Chemistry 58 1839 (1986) Nevinnaya, L.V. and Klyachko, Y.A. Soviet Journal of Water Chemical Technology 10 71 (1988) Wee, V.T and Kennedy, J.M. Analytical Chemistry 54 1631 (1982) Basson, W.D. and Van Staden, J.F. Analyst (London) 103 998 (1978) Skully, F.E., Oglesby, D.M. and Buck, H.J. Analytical Chemistry 56 1449 (1984) Van de Haar, G., Pijper-Noordhoff, F.M. and Strikwerda, K. H2 12 420 O (1972) Eremin, Yu.G. and Kopylova, G.A. Zavod. Lab. 89 1065 (1973) Nehring, H., Pharmazie 27 741 (1972) McCarthy, J.J. Limnology and Oceanography 15 309 (1970) Emmet, R.T. Analytical Chemistry 41 1648 (1969) Rogers, D.S. and Pool, K. Analytical Letters (London) 6 801 (1973) Chu, J.P., Kirsch, E.J. and Born, G.S. Bulletin of Environmental Contamination and Toxicology 6 343 (1971)
476
Nitrogen-containing compounds
168 Godjehann, M., Preiss, A., Mugge, C. and Wünsch, G. Analytical Chemistry 69 3832 (1977) 169 Tompkins, B.A. and Griest, W.H. Analytical Chemistry 68 2533 (1996) 170 Tompkins, B.A., Griest, W.H. and Higgins, C.E. Analytical Chemistry 67 4387 (1995) 171 Kari, F.G. Dissertation. ET H. No. 10698. Dübendorfer Switzerland (1994) 172 Schürch, S. and Dubendorfer, G. Mitt. Geb. Lebensmittelunters 80 324 (1989) 173 Bergers, P.J.M. and de Groat, A.C. Water Research 28 639 (1994) 174 Sillanpaa, M., Kokkonen, B. and Sihvonen, M.L. Anal. Chim. Acta 303 187 (1995) 175 Ye, L. and Lucy, C.A. Analytical Chemistry 67 2534 (1995) 176 Nowack, A., Kari, F.G., Hilger, S.U. and Sigg, L. Analytical Chemistry 68 561 (1996) 177 Petty, R.L., Michel, W.C., Snow, J.P. and Johnson, K.S. Anal. Chim. Acta 142 299 (1982)
Chapter 7
Phosphorus-containing compounds
7.1 Alkyl and aryl phosphates 7.1.1 Natural waters 7.1.1.1 Gas chromatography Murray [1] has described a gas chromatographic method for the determination in water of triarylphosphate esters (1mol S-140, tricresyl phosphate, cresol phosphate). These substances are used commercially as lubricant oil and plastic additives, hydraulic fluids and plasticizers. The method involves extraction from the samples, hydrolysis and measurement of the individual phenols by gas chromatography as the trimethylsilyl derivatives. The lower detection limit was about 3ppm. In this procedure a weighed sample was placed in a 20ml ampoule and 10ml of 5% potassium hydroxide in 95% methanol added. The ampoule was sealed and autoclaved at 25psi (172kPa) for 90min. When cool, the ampoule was opened and the contents washed into 950ml of distilled water, acidified with about 5ml 6M hydrochloric acid to pH1–2 and made up to 1000ml. This was transferred to a separately funnel, 5µL of o-xylene were added by syringe as an internal standard, and the mixture was extracted once with 50ml of chloroform. The solvent layer was evaporated to 1–2ml and treated with Tri-Sil concentrate to form trimethylsilyl derivatives. This was allowed to react overnight and analysed by gas chromatography the following day. The concentrations of the individual phenols were calculated from calibration graphs and the composition of the ester was determined. A gas chromatograph with flame ionization detectors was used for the analysis and the column used was 8ft × 1/8 in (2.4m × 3mm) stainless steel packed with 5% Imol on Chromosorb W, AW and SCMS treated 80–100 mesh. A temperature programme from 80 to 120°C at 4°C min-1 was used.
478
Phosphorus-containing compounds
Ishikawa et al. [2] have developed procedures for determining these compounds in river waters. The procedure involves extraction of the sample with dichloromethane followed by gas chromatography using a flame photometric detector and gas chromatography–mass spectrometry after clean-up through a Florasil column. 7.1.2 Estuary and sea waters 7.1.2.1 X-ray fluorescence spectrometry Ahern et al. [3] have observed that organophosphorous compounds are coprecipitated with cobalt and pyrrolidine dithiocarbonate and suggest that this might be a suitable means of preconcentrating the phosphorous fraction of harbour water prior to analysis by X-ray fluorescence spectrometry. 7.1.3 Potable waters 7.1.3.1 Miscellaneous Williams and Le Bel [4] examined potable waters for 29 cities in Canada, for tri(haloalkyl), trialkyl and triaryl phosphate. A comparison of triaryl phosphate levels with raw water sources indicated that higher levels were found in potable water from rivers than lakes and ground waters have very low levels. The three central drainage basins of the St Lawrence River, Great Lakes and Lake Winnipeg had relatively high levels of triaryl phosphate. The fraction of triaryl phosphate values over 10ng L-1 was very small, but about 75% of the tributoxyethyl phosphate values were over l0ng L-1 possibly through contamination from the rubber gasket industry. Trace levels of triaryl phosphate appear to be present in Canadian potable water supplies sampled at the 29 water treatment plants. 7.2 Adenosine triphosphate An estimate of the quantity of living microorganisms (biomass) in an aquatic environment can be a useful tool to assess water quality. Both phytoplankton and microzooplankton biomass can be estimated by direct microscopic determination of volume and number. Other microorganisms (for example, bacteria) have been estimated by various plating techniques (colony counting) and extinction–dilution techniques. The techniques are often time-consuming, expensive and subject to inherent sources of error. Some of the major problems include the following. Direct microscopic counting of bacteria may yield higher estimates because of the difficulty in distinguishing bacteria from bacteria-sized
Adenosine triphosphate
479
inert particles. There is also the inability to differentiate between viable and non-viable cells, and the problems of cell aggregation, for bacteria and algae. Low estimates may be produced by plating and extinction techniques which are selective because of the chemical composition of the media and inherent physical parameters such as temperature and pressure. The extinction–dilution technique may also be biologically selective in that only types capable of growth in specialized media will grow to measurable size. For a total biomass determination, it is desirable to measure some biochemical that is present in all living cells but is not associated with non-living particulate material. This cellular constituent must have a short survival time after death, so that it would be specific for viable biomass. It must likewise be present in proportion to some measure of the total biomass for all microorganisms – algae, bacteria, fungi and protozoans. A biochemical which seems to meet these requirements is adenosine triphosphate. Adenosine triphosphate is the primary energy donor in cellular life processes. Its central role and biological and chemical stability make it an excellent indicator of the presence of living material. The level of endogenous adenosine triphosphate, which is the amount of adenosine triphosphate per unit biomass bacteria, algae and zooplankton is relatively constant when compared to the cellular organic carbon content in several species and throughout all phases of the growth cycle. In studies where cell viability was determined the concentration of adenosine triphosphate per viable cell remains relatively constant during periods of starvation. The quantity of adenosine triphosphate, therefore, can be used to estimate total living biomass. 7.2.1 Natural waters 7.2.1.1 Luciferin–luciferase bioluminescence technique Tobin et al. [5] give details of two extraction procedures for the determination of adenosine triphosphate in environmental samples by luciferin–luciferase assay. Both methods are suitable for natural waters Shoaf and Lium [6] have also discussed this technique. Andre et al. [7] discuss the determination of adenosine triphosphate by luciferin–luciferase assay. This method was applied to the determination of adenosine triphosphate in bacterial colonies filtered from samples of polluted water after incubation for different periods. The adenosine triphosphate was extracted from the residue in the filter and the amount compared with the biochemical oxygen demand of the filtered water. The oxygen uptake rate and the rate of formation of adenosine triphosphate were then plotted against time, the two curves being similar up to 3–4
480
Phosphorus-containing compounds
days’ incubation, after which adenosine triphosphate production declined markedly, although oxygen uptake continued to increase. 7.2.1.2 Miscellaneous Siegrist [8] has described a method for the determination of adenosine triphosphate in lake water. The procedure can be applied in routine limnological work by freezing the filter residue samples in liquid nitrogen for transport to the laboratory for analysis. Sample preservation between sampling and analysis is a problem [6]. Quick freezing of the sample in an acetone dry-ice bath maintains the adenosine triphosphate concentration in the sample at a constant level. 7.2.2 Waste waters 7.2.2.1 Luciferin–luciferase bioluminescence technique Hysert et al. [9–11] have applied a bioluminescence adenosine triphosphate assay method to brewery waste water studies. They describe an assay using partially purified luciferase and synthetic firefly luciferin for the bioluminescence reaction, a liquid scintillation counter in the out of coincidence phase and a sludge adenosine triphosphate technique involving dimethyl sulphoxide at room temperature. Experiments with several pure bacteria cultures showed good linear correlation between adenosine triphosphate and mixed liquor suspended solids, return sludge suspended solids and effluent suspended solids. When adenosine-51-diphosphate and adenosine-51-monophosphate as well as adenosine triphosphate assays were to be performed, the extractions were done with boiling acetone [11–13]. Typically 0.1ml sample was added to 1ml boiling acetone in a 7ml vial immersed in a 90°C water bath and the acetone was removed by blowing a stream of air over the liquid, 10µl of extracts were added to 1.0ml, pH7.0, 10mM tris(hydroxymethyl)aminomethane (TRIS)–3.5mM magnesium sulphate buffer and 10µl of a purified luciferase synthetic D-luciferin premix were added to initiate the bioluminescence reaction. The light output of the reaction was measured with a scintillation counter. 7.2.3 Sewage effluents 7.2.3.1 Luciferin–luciferase bioluminescence technique Patterson et al. [14] carried out determinations of adenosine triphosphate in activated sludge. The method involved the use of firefly lantern extract.
Inositol phosphate esters
481
Hysert et al. [9] demonstrated that interference and inhibition of the bioluminescence assay by extract components was negligible by a standard adenosine triphosphate method [15]. The reproducibility of the overall method for determining the adenosine triphosphate content of activated sludge, which includes sampling, dimethyl sulphoxide extraction and adenosine triphosphate assay, was considerably poorer than that observed for the adenosine triphosphate assay alone or for adenosine triphosphate assays of pure cultures. The relative standard deviations for the latter assays were 2–4%, whereas those for the activatedsludge determinations were in the 7–11% range. This greater method of variance no doubt results from difficulties of reproducibly sampling the heterogenous, clumped biological flocs that constitute activated sludge. 7.2.4 Trade effluents 7.2.4.1 Luciferin–luciferase bioluminescence technique This technique has been applied to trade effluents [9, 16]. Kucnerowicz and Verstraete [16] carried out direct measurements of micobial adenosine triphosphate in activated-sludge samples. The method uses an activated-sludge apparatus designed for determining the biodegradability of anionic detergents. Mixed liquors are diluted with triethanolamine buffer, homogenized, mixed with adenosine triphosphate releasing agent and the luminescence of the mixture is measured after addition of luciferase. Results obtained were in agreement with literature data for adenosine triphosphate in activated sludge. Relationships established between adenosine triphosphate content of sludges and other sludge parameters indicate that adenosine triphosphate determination could be used as a method of monitoring activated-sludge treatment processes. 7.3 Inositol phosphate esters 7.3.1 Natural waters 7.3.1.1 Miscellaneous Weimer and Armstrong [17] have described methods for the determination of inositol phosphate.
482
Phosphorus-containing compounds
7.4 Organophosphorus compounds – general 7.4.1 Natural waters 7.4.1.1 Gas chromatographic detection with supported copper– cuprous oxide island film Organophosphorus compounds have been detected in environmental waters at the parts per million level using a supported copper–cuprous oxide island film gas chromatographic detector. Both alternating current [18] and direct current studies were used [19]. 7.4.1.2 Conducted chemiresistant sensors for gas chromatographic detection Grate et al. [20] have studied the role of selective sorption in chemiresistant sensors for the gas chromatographic detection of Organophosphorus compounds. 7.4.1.3 Surface acoustic wave sensors for gas chromatographic detection Grate et al. [21] have described a smart sensor system for detecting traces of Organophosphorus compounds. Vapour detection of these compounds is achieved employing a temperature controlled array of surface acoustic wave detectors, automated sample preconcentrations and pattern recognition. Down to 0.01mg m-3 of Organophosphorus compounds can be detected on the detector surface. 7.4.1.4 Miscellaneous Verweij et al. [22] studied the hydrolysis of some methylphosphonites and methylphosphonates in the pH range 4.5–7.5 by phosphorus-31 nuclear magnetic resonance spectrometry. For methylphosphonites, a two-step reaction mechanism was observed in which formation of the corresponding methylphosphonates was followed by production of methylphosphrnic acid. Implications of the results obtained on the applicability of a verification procedure for nerve gases in surface waters are discussed. References 1 2
Murray, D.A.J. Journal of Fisheries Research Board, Canada 32 457 (1975) Ishikawa, S., Taketomi, M. and Shinokara, R. Water Research 19 119 (1985)
References
3 4 5 6 7 8 9 10 11 12
13
14 15 16 17 18 19 20 21 22
483
Ahern, F, Eckhert, J.M., Hain, S.F., Leggett, K.E.A., Payne, N.C. and Williams, K.L. Anal. Chim. Acta 199 259 (1987) Williams, D.T., Le Bel, G.L. Bull. Environmental Contamination and Toxicology 27 450 (1981) Tobin, R.S., Ryan, J.F. and Afghan, B.K. Water Research 12 783 (1978) Shoaf, W.T. and Lium, B. Journal of Research of the U.S. Geological Survey 4 241 (1976) Andre, M., Van Beneden, P. and Bassleer J. Tribune du Cebedeau 31 251 (1978) Siegrist, L.H. Schweizerische Zeitschrift für Hydrologie 38 49 (1976) Hysert, D.W., Knudson, F.B., Morrison, M.N., Van Gheluwe, G. and Lom, T. Biotechnol. Bioengng 21 1301 (1979) Hysert, D.W., Kovecses, F. and Morrison, N.M. J. Am. Soc. Brew. Chem. 34 145 (1976) Hysert, D.W. and Morrison N.M. J. Am. Soc. Brew. Chem. 35 160 (1970) Chappelle, E.W., Picciolo G.L., Curtis, C.A., Knust, E.A., Nibley, D.A. and Vance, R.B. Laboratory Procedures Manual for the Firefly Luciferase Assay for Adenosine Triphosphate (ATP). Goddard Space Flight Centre, NASA, TMX70926 (1975) Knust, E.A., Chappelle, E.W. and Picciolo, G.L. In Analytical Applications of Bioluminescence and Chemiluminescence, Chappelle, E.W. and Picciolo, G.L (eds), NASA Scientific and Technical Information Office, Washington DC, p. 27 (1975) Patterson, J.W., Brezonick, P.L. and Putnam, H.D. Environmental Science and Technology 4 569 (1970) Lundin, A. and Thore, A. Analytical Biochemistry 66 47 (1975) Kucnerowicz, F. and Verstraete, W. Journal of Chemical Technol Biotechnol 29 707 (1979) Weimer, W.C. and Armstrong, D.E. Anal. Chim. Acta 94 35 (1977) Kolesar, E.S. and Walser, R.M. Analytical Chemistry 60 1737 (1988) Kolesar, E.S. and Walser, R.M. Analytical Chemistry 60 1731 (1988) Grate, J.W., Klusty, M., Borger, W.R. and Snow, A.W. Analytical Chemistry 62 1927 (1990) Grate, J.W., Rose Pehrsson, S.L., Venezky, D.L., Klusty, M. and Wohitjen, H. Analytical Chemistry 65 1868 (1993) Verweij, A., Dekker, W.H., Beck, H.C. and Boter, H.L. Anal. Chim. Acta 151 221 (1983)
Chapter 8
Sulphur-containing compounds
8.1 Mercaptans and disulphides 8.1.1 Natural waters 8.1.1.1 Titration method Duane and Stock [1] have discussed a titration method for determining thiols. 8.1.1.2 Gas chromatography Leek and Bagander [2] determined reduced sulphur compounds in natural waters by gas chromatography using a flame ionization detector. Substances determined include methyl mercaptan, dimethyl sulphide, hydrogen sulphide and carbon disulphide. Detection limits ranged from 0.2mg L-1 (carbon disulphide) to 0.6ng L-1 (methyl mercaptan). Heuatsch and Jutter [3] have discussed the gas chromatographic determination of dimethyl sulphide in lake water. 8.1.2 Sea water 8.1.2.1 Gas chromatography The procedure discussed above in Section 8.1.1.1 has also been applied to sea water. 8.1.3 Waste waters 8.1.3.1 Gas chromatography Vitenberg et al. [4] have described a gas chromatographic method for the determination of traces (down to 10-6–10-7%) of sulphur compounds, such
Mercaptans and disulphides
485
as hydrogen sulphide, mercaptans, sulphides and disulphides, in industrial waste waters (kraft paper mill effluents) by a combination of headspace analysis and microcoulometry. This method, described below, increases the analytical sensitivity 102–103 times without any preliminary concentration of the sample. A sample of the solution to be analysed of volume V without the gas L phase is drawn into the variable-volume device [5]. A 1:1 KCl–HCl buffer solution of pH2 containing 14% of sodium sulphate is added to the sample through the elastic rubber plug with the help of a hypodermic syringe. After this, air of volume V (an inert gas is preferable for g prolonged equilibration) is drawn into the vessel with the solution to be analysed and the system is maintained for 30min at a constant temperature with periodic shaking. After equilibration, the concentration of the sulphur compounds in the gas phase above the solution is determined. To do this, the equilibrium gas is displaced by the plunger from the vessel, filling the same loop of the gas sample valve which is used to inject the analyte into the chromatographic column. On measuring the areas and heights of the corresponding peaks on the chromatogram I , one determines the concentration of each compound of s interest in the gas phase from the calibration data I = f( g) available for a s C given sample loop. The content of each sulphur compound in the solution is calculated by the equation below, taking into account dilution by the buffer solution (pH2) containing 7% sodium sulphate,
where K is the partition coefficient and is concentration of microimpurity in the original solution. At a concentration level of 1 ppm sulphur compounds, the analytical error does not exceed 8% for the flame ionization detector and 12% for the microcoulometric detector, of the given amount of the compound. When analysing solutions with concentrations of not more than 0.lppm, the analytical error in the region of the highest sensitivity reaches 15% and 20% for the flame ionization and the microcoulometric detector, respectively.
486
Sulphur-containing compounds
8.1.4 Trade effluents 8.1.4.1 Gas chromatography Devonald et al. [6] have described a gas chromatographic method utilizing a flame photometric detector for the determination of hydrogen sulphide, sulphur dioxide, methane diol, dimethyl sulphide and dimethyl disulphide in industrial effluents in amounts down to 0.1mg L-1. Sulphurfree organic compounds often present in these effluents, such as lactones, alcohols and terpenes, do not interfere. Vitenberg et al. [4, 5] have described a gas chromatographic method for the determination of traces down to micrograms per litre sulphur compounds, such as hydrogen sulphide, mercaptans, sulphides and disulphides in industrial waste waters, by a combination of headspace analysis with microcoulometric detector and flame ionization detectors. The headspace sampling technique increases the analytical sensitivity by 102–103. 8.1.5 Sewage effluents 8.1.5.1 Gas chromatography Bailey and Viney [7] have applied gas chromatography to the investigation of odours produced at sewage treatment plants. Samples of ambient air at a sewage treatment works were taken using Tedlar bags and also by using traps containing Tenax GC on site. Their studies showed that methyl mercaptan found in the headspace of these samples was produced by aerobic decomposition of hexyl mercaptan. The method described by Jenkins et al. [8] involves gas chromatography coupled with the sulphur-specific flame photometric detector which can detect 0.25ng hydrogen sulphide and 0.5ng of methyl mercaptan. The method has been used to trace sources of odour due to mercaptans in the air surrounding a sewage works and in waste waters and sewer gases. They used these techniques to study the fate of various organosulphur compounds in raw sewage and activated sludge. Jenkins et al. [8] used a gas chromatograph equipped with a Melpar flame photometric detector and a Perkin–Elmer 3920B gas chromatograph equipped with a linearized Perkin–Elmer flame photometric detector. Each instrument also had a flame ionization detector. An FEP Teflon column (18in × 0.625in i.d. (0.46m × 15mm)) containing acetone-treated Poropak QS was used for all analyses. The PE 3920B used a glass-lined vaporizer injector and the interface lines were
Hydromethanesulphinite salts
487
in stainless steel. For the Tracer 550, the Poropak QS column was connected to glass capillary tubing which was inserted into the injector and outlet barrels. Teflon-backed septa were used in the injectors of both instruments. 8.1.5.2 Gas chromatography–mass spectrometry Konig et al. [9], using gas chromatography–mass spectrometry, have identified dimethyl di-, tri- and tetrasulphides in sewage works samples. These substances can be used by microorganisms as precursors of sulphuric acid, which would explain unexpectedly high corrosion rates of concrete pipes. 8.1.5.3 Miscellaneous Roe [10] has discussed the determination of volatile compounds (methanoldiol, dimethyl sulphide) and also hydrogen sulphide and sulphur dioxide in sewage samples. 8.2 Benethonium salts 8.2.1 Waste waters 8.2.1.1 Spectrophotometric method These compounds have been determined in waste water by a spectrophotometric method based on reaction with tetrabromophenolphthalein ester [11]. 8.3 Hydromethanesulphinite salts 8.3.1 Trade effluents 8.3.1.1 Polarography These substances and their decomposition products (formaldehyde, zinc and sulphites) have been determined by polarographic procedure in waste waters [12] originating from textile industries. In an alkaline medium it is possible to have well-defined and separated waves relative to the oxidation of hydroxymethanesulphinite ion and to the reduction of formaldehyde and zincate ion. The quantitative determination of these compounds is carried out with standard addition methods, since the
488
Sulphur-containing compounds
concentration–current relationship is linear. The sulphite is determined in 0.1M perchloric acid and in the same sample it is possible to determine hydroxymethanesulphinite salts, formaldehyde and zinc after addition of sodium hydroxide up to 0.1M. An indirect method for the determination of hydroxymethanesulphinite salts is described. This method is based on the determination of formaldehyde released by means of a weak oxidation of these salts which is stoichiometrically proportional to total hydroxymethanesulphinite content. 8.4 Benzothiazole, 2-mercaptobenzothiazole and 2-(4-morpholinyl)benzothiazole 8.4.1 Trade effluents 8.4.1.1 Gas chromatography–mass spectrometry Jungclaus et al. [13] used gas chromatography–mass spectrometry to identify benzothiazole and 2-mercaptobenzothiazole in tyre manufacturing plant effluents. The gas chromatograph was equipped with a flame ionization detector. Separations were achieved on a 180cm × 0.32cm stainless steel column packed with 3% SP 2100 (methyl silicone fluid) on 80–100 mesh Supelcoport and was temperature programmed for 70–300°C at 16°C min-1. Approximately 44 compounds were identified in the waste water samples including benzothiazole and 2-mercaptobenzothiazole at concentrations, respectively, of 0.06 and 0.03µg L-1. Kumata et al. [24] have described a procedure based on gas chromatography with a flame photometric detection for the determination of 2-(4-morpholinyl)benzothiazole in environmental water samples including river waters and street run-off waters. 2-(4morpholinyl)-benzothiazole exists in automobile tyre rubber as an impurity of a vulcanization accelerator and has been proposed as a potential molecular marker of street run-off [25]. Kumata et al. [24] describe an analytical method for 2-(4-morpholinyl)benzothiazole in environmental samples (e.g., street dusts and river sediments) by gas chromatography. The method relies upon extraction with a 6:4 (v/v) mixture of benzene and methanol, purification by acid extraction, and adsorption column chromatography, followed by determination using capillary gas chromatography equipped with a sulphur-selective detector (i.e. FPD). The recovery of 2-(4-morpholinyl)benzothiazole for the entire procedure was 85%, and the relative standard deviation for four replicated analyses was 1.5%. The detection limit was 0.08ng injected 2-(4-morpholinyl)-benzothiazole, corresponding to 0.20ng g-1 of
Dimethyl sulphoxide
489
dry sample. The selectivity and sensitivity of this method permit the determination of 2-(4-morpholinyl)benzothiazole at trace levels (e.g., ng g-1) encountered in environmental samples. 8.4.1.2 High-performance liquid chromatography Cox [14] determined mercaptobenthiazole in trade effluent samples by a high-performance liquid chromatographic procedure. An ultraviolet detector operating at 325nm was used to monitor the column effluent. The column was constructed from stainless steel tubing (15cm × 4.6mm) and was packed with Microkosorb Sil60 (5µm silica gel) at 3500psi (24.1MPa) from a slurry in 2,2,4-trimethylpentane. The packing material was retained in the column by stainless steel wire mesh of nominal pore size 8µm (Sankey Wire Weaving, Warrington, UK) inserted into a drilledout Swagelok coupling. A similar disc of wire mesh was pressed on the top of the column packing and was retained by a plug of silanized glass wool. Ethanol–2,2,4-trimethylpentane (1:9) was used as the mobile phase with flow rate of 1mL min-1. All solvents used were of spectroscopic quality (Fisons, Loughborough, UK). The aqueous sample (2mL) was acidified with two drops of concentrated hydrochloric acid. This mixture was shaken with chloroform (2mL) for 1min using a flask shaker. Aliquots (2µL) of the chloroform layer were used for the chromatographic analysis. 8.5 Dimethyl sulphoxide 8.5.1 Natural waters Dimethyl sulphoxide is a well-known chemical used as a solvent, a lubricant and an additive in industrial applications, but its occurrence in aquatic environments arises mainly from natural processes. In fact, dimethyl sulphoxide is a major pool of dissolved sulphur in sea water, where it occurs at nanomolar levels, and is a significant constituent in other natural waters and in the atmosphere. The implication of dimethyl sulphoxide in the biogeochemical cycle of dimethyl sulphide has been anticipated. Dimethyl sulphide is the most abundant volatile sulphur compound in surface sea water, and a key species of the global sulphur cycle. Sea water dimethyl sulphide is produced by plankton, mostly from algal dimethyl sulphoniopropionate. Upon release to the air, dimethyl sulphide undergoes oxidation to form acidic aerosols and condensation nuclei, thereby affecting tropospheric chemistry and cloud reflectiveness over the oceans. Current evidence suggests that the dynamics of dimethyl sulphide in water is influenced
490
Sulphur-containing compounds
by photochemical and microbial processes. Laboratory studies have shown that dimethyl sulphoxide can be a potential source and sink for dimethyl sulphide through these two types of processes. However, the significance of the role of dimethyl sulphoxide in the dimethyl sulphide cycling still has to be assessed. 8.5.1.1 Gas chromatography Simo et al. [26] have discussed the determination of nanomolar concentrations of dimethyl sulphoxide, along with dimethyl sulphide and dimethyl sulphoniopropionate, at nanomolar levels in natural waters. After removal of dimethyl sulphide by purge and cryotrapping, dimethyl sulphoniopropionate is removed by the same method after alkaline hydrolysis, and dimethyl sulphoxide is reduced to dimethyl sulphide using a combination of sodium borohydride and hydrochloric acid. The dimethyl sulphide produced is stripped, cryotrapped and analysed by gas chromatography. Detection of 3pmol of dimethyl sulphoxide was achieved, resulting in a detection limit of 0.05nM for a 50mL sample. Mean yield for standards in the range of 0.7–130nM (n = 31) was 95%, and mean precision (as coefficient of variation) was 14%. Precision for replicates of natural sea water samples was always =10%. Mean yields of the sequential analysis of dimethyl sulphide plus dimethyl sulphoxide plus dimethyl sulphoniopropionate (1.5–25nM) standard mixtures in sea water were >90% for the three species. Filtered sea water samples stored frozen (-20°C) showed no significant changes in dimethyl sulphoxide concentration. Since dimethyl sulphoniopropionate is the only compound, other than dimethyl sulphoxide that gives rise to dimethyl sulphide upon reduction with sodium borohydride, tests were performed to ensure that dimethyl sulphoniopropionate is quantitatively removed before the dimethyl sulphoxide analysis. Boranes were identified as the substances that produced two major peaks in the chromatogram of the reduction products. Adequate chromatographic conditions were established to avoid their coelution with dimethyl sulphide. This entire protocol allows the sequential determination of dimethyl sulphide, dimethyl sulphoxide plus dimethyl sulphoniopropionate in natural waters and is suitable for field work, so it should be very useful in studies of dimethyl sulphur biogeochemistry. Andreae [15] has described a gas chromatographic method for the determination of nanogram quantities of dimethyl sulphoxide in natural waters, sea water and phytoplankton culture waters. The method involves a chemical reduction to dimethyl sulphide, which is then determined gas chromatographically using a flame photometric detector. Andreae [15] investigated two different apparatus configurations. One consisted of a reaction/trapping apparatus connected by a six-way
Dimethyl sulphoxide
491
valve to a gas chromatograph equipped with a flame ionization detector; the other apparatus combined the trapping and separation functions in one column, which was attached to a flame photometric detector. The gas chromatographic flame ionization detector system was identical to that described by Andreae [15] for analysis of methylarsenicals, with the exception that a reaction vessel which allowed the injection of solid sodium borohydride pellets was used. The flame photometric system (Fig. 8.1) is modified after a design by Braman et al. [16]. The first stage in both systems is a reduction vessel, which contains the sample and buffer solutions. Helium is bubbled through the solution by a glass diffuser. A side port allows the injection of the reducing solutions by a hypodermic syringe through a Teflon-coated silicone septum attached by a Teflon Swagelok fitting. In a modified design, a short, bent piece of glass tubing which can hold a borohydride pellet is attached to the upper part of the reaction vessel by a ground glass joint. By turning this tubing, the pellet can be dropped into the solution without opening the system to the atmosphere. For the borohydride reduction, 0.1mL concentrated hydrochloric acid is added per 25mL sample; for the reduction with chromium (II) chloride, 2.5mL of acid are added. The reaction vessel is then attached to the system and purged for 5min with the helium stream to remove air and volatile sulphur compounds. Scrubbing is then continued until a zero blank is obtained.
Fig. 8.1 Apparatus for the reduction of DMSO to DMS and the flame photometric detection of DMS (FPD system). (Reprinted with permission from Andreae [15]. Copyright (1980) American Chemical Society.)
492
Sulphur-containing compounds
After the purging period, the cold trap is immersed in liquid nitrogen, and the reductant (chromium (II) chloride, sodium borohydride pellet or solution) is injected. When sodium borohydride solution is used, it has to be slowly injected into the reactor (ca. 1min per 2mL) to avoid an excessively fast reaction. The reductant amounts used are: sodium borohydride, one 0.25g pellet or 2mL of 4% solution; chromium (II) chloride, 5mL of a 2M solution. The reaction times are 6 and 30min for the borohydride and chromium (II) reductions, respectively. The helium stream is continuously purging the solution during this period in order to remove the volatile dimethyl sulphide formed by reduction of dimethyl sulphoxide from the solution. The gas stream is dried by passing through a U-tube at –30°C which removes all of the water and none of the dimethyl sulphide. The dimethyl sulphide and other reaction products in the gas stream are collected in a cold trap immersed in liquid nitrogen. For the chromatographic–flame ionization system, this trap is filled with silanized glass wool. After the collection time, the trap is switched into the carrier gas stream of the gas chromatograph by the six-way valve, then heated rapidly by immersing in hot water. The dimethyl sulphide is then separated on the column from other reaction products and detected by the flame ionization detector. In the flame photometric system, the trap serves both to collect the reaction products and to achieve the gas chromatographic separation. For this purpose, a trap filled with a gas chromatographic packing is used. After the reaction time, the liquid nitrogen is removed, the variable transformer is switched to provide 7V to the heating coil. The dimethyl sulphide elutes in a sharp peak after 1.1min and enters the flame photometric detector. Using 100mL samples, a detection limit of 0.01µg L-1 of dimethyl sulphoxide was achieved by these procedures. The flame ionization detector shows a completely linear response in the lower range with a decrease in slope starting at the 100ng level. 8.5.1.2 Miscellaneous Andreae [15] showed that dimethyl sulphoxide is a common constituent in natural waters. Its occurrence in sea water is restricted to the zone of light penetration. This fact and the abundance of dimethyl sulphoxide in the medium after the growth of photoplankton suggests that it occurs as an end product of algal metabolism. The occurrence of dimethyl sulphoxide in rain may be due either to the release of this substance from the biosphere into the atmosphere or to the atmospheric oxidation of dimethyl sulphide to dimethyl sulphoxide.
Chlorolignosulphonic acids
493
8.6 Calcium lignosulphate 8.6.1 Trade effluents 8.6.1.1 Gel permeation chromatography Sagfors and Starck [17] used gel permeation chromatography to study substances of high molecular mass in acid and alkaline kraft pulp bleaching effluents. 8.6.1.2 Ultraviolet spectroscopy This substance has been determined by ultraviolet spectroscopy in sulphite-containing effluents [18]. 8.7 Thiafluoron 8.7.1 Potable waters 8.7.1.1 High-performance chromatography This technique has been applied to the determination of thiafluoron in potable water [19]. Separation was achieved on a C Sep Pak column and 18 a fluorescence and electrochemical detector were used to determine microgram per litre quantities of this compound. 8.8 Chlorolignosulphonic acids 8.8.1 Natural waters 8.8.1.1 Pyrolysis–gas chromatography–mass spectrometry with single-ion monitoring Van Loon et al. [20] have used this technique to determine Chlorolignosulphonic acids of molecular weight greater than 1000 in natural waters in amounts down to 0.1µg L-1. Between 18 and 310µg L-1 of this substance were present in River Rhine water.
494
Sulphur-containing compounds
8.8.2 Potable waters 8.8.2.1 Pyrolysis–gas chromatography The pyrolysis–gas chromatography technique [20] discussed in Section 8.8.1.1 has been applied to potable waters. Up to 60µg L-1 were found in typical potable water samples. 8.8.3 Trade effluents 8.8.3.1 Pyrolysis–gas chromatography The pyrolysis–gas chromatography technique [20] discussed in Section 8.8.1.1 has been applied to sulphite pulp mill effluents. 8.9 Tetrahydrothiophen 8.9.1 Natural waters 8.9.1.1 Gas chromatography The determination of tetrahydrothiophen has been discussed by Garlucci et al. [22]. 8.10 Organosulphur compounds, general Grate et al. [21] have discussed the smart sensor system for detecting down to 0.5mg M-1 organosulphur compounds in environmental waters. This consisted of a temperature controlled array of surface acoustic wave sensors (SAWS), with automated sample preparation and pattern recognition. Organosulphur compounds have been preconcentrated on XAD-2 and XAD-4 macroreticular resins prior to solvent desorption and analysis by headspace gas chromatography [23]. References 1 2 3 4
Duane, L. and Stock, J.T. Analytical Chemistry 50 1891 (1978) Leek, C. and Bagander, L.E. Analytical Chemistry 60 1680 (1988) Heuatsch, J.J. and Jutter, F. Journal of Chromatography 445 97 (1988) Vitenberg, A.G., Kuznetsova, L.H., Butaeva, I.L. and Ishakov, M.D. Analytical Chemistry 49 128 (1977)
References
5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26
495
Vitenberg, A.G., Butaeva, I.L. and Dimitova, Z.St. Chromatographia 8 693 (1975) Devonald, R.H., Serenins, R.S. and McIntyre, A.D. Pulp Paper Magazine, Canada 73 50 (1972) Bailey, J.C. and Viney, N.J. Gas chromatographic investigation of odour and a sewage treatment plant. Water Research Centre, Medmenham Laboratory, Medmenham, UK. Technical Report No. TR 125 December (1979) Jenkins, L.L., Gute, J.P., Krasner, S.W. and Baird, R.B. Water Research 14 441 (1980) Konig, W.A., Ludwig, K., Rinken, M., Stalting, K.H. and Gunther, W. Journal of High Resolution Chromatography. Chromatography Communications 3 415 (1980) Roe, A.B. Journal of the Institute of Water Engineers and Scientists 36 118 (1982) Tsubauchi, M. Bull. Chem. Soc. Japan 44 1560 (1971) Piccardi, E.B. and Cellini, P. Water, Air and Soil Pollution 9 301 (1978) Jungclaus, G.A., Game, L.H. and Hites, R.A. Analytical Chemistry 48 1894 (1976) Cox, G.B. J. Chromatography 116 244 (1976) Andreae, M.O. Analytical Chemistry 52 150 (1980) Braman, R.S., Ammons, J.M. and Bricker, J.L. Analytical Chemistry 50 992 (1978) Sagfors, P.E. and Starck, B. Water Science Technology 20 49 (1988) Bau, S., Glauser-Soljam, M. and Smarlagic, M. Biotechnology Bioengineering 21 1917 (1979) Putzien, J.Z. Wasser Abwasser Forsch. 19 228 (1986) Van Loon, W.M.G.M., Bron, J.S. and de Groot, B. Analytical Chemistry 65 1726 (1993) Grate, J.W., Pehrsson, S.L., Venezky, D.L., Klusy, M. and Wohitjen, H. Analytical Chemistry 65 1868 (1993) Garlucci, G., Airoldi, L. and Farelli, R. Journal of Chromatography 287 425 (1984) Przazny, A.J. Chromatography 346 61 (1988) Kumata, H., Takada, H. and Ogura, N. Analytical Chemistry 68 1976 (1996) Spies, R.B., Andresen, B.D. and Rice, D.W. Nature ( London ) 327 697 (1987) Simo, R., Grimalt, J.O. and Albarges, J. Analytical Chemistry 68 1493 (1996)
Chapter 9
Insecticides and herbicides
9.1 Chlorinated insecticides 9.1.1 Natural waters 9.1.1.1 Gas chromatography Work on the determination of chlorinated insecticides has been almost exclusively in the area of gas chromatography using different types of detection systems, although a limited amount of work has been carried out using liquid chromatography and thin-layer chromatography. By its nature, gas chromatography is able to handle the analysis of complex mixtures of chlorinated insecticides. It is not surprising therefore, that much of the published work discussed in this section is concerned with the analyses of mixtures of different types of chlorinated insecticides as found in environmental samples. Work on the determination of individual insecticides is reported at the end of this section. Work on the determination of polychlorinated biphenyls (PCBs) and mixtures of these with chlorinated insecticides is reported in Section 9.2 on PCB and PCB–organochlorine pesticide mixtures. Various solvents have been used for the preliminary separation of chlorinated insecticides from natural waters prior to gas chromatographic analysis. The most commonly used solvents include hexane, petroleum ether, mixtures of hexane and benzene or toluene, benzene, diethyl ether, methylene dichloride and acetonitrile (Table 9.1). The American Public Health Association [6, 29] published an early gas chromatographic method for the solvent extraction and gas chromatographic determination of 11 chlorinated insecticides in water samples in amounts down to 0.005mg L-1, p,p’-DDE, p,p’-DDT, Aldrin, Dieldrin, Endrin, Heptachlor, Heptachlor epoxide, Lindane, Isodrin and Methoxychlor. The insecticides Carbophenothion, Chlordane, Dioxathion, Diazinon, Ethion, Malathion, Parathion methyl, methyl
Chlorinated insecticides
497
Table 9.1 Extraction solvents for concentration of chlorinated insecticides prior to gas chromatographic analysis
498
Insecticides and herbicides
Table 9.1 Continued
Chlorinated insecticides
499
Trithion, Parathion, Toxaphene and VC-13 may be determined when present at higher levels. Also, the chemicals Chlordene, hexachlorobicycloheptadiene and hexachlorocyclopentadiene, which are pesticide manufacturing precursors, may be analysed by this method. The insecticides are extracted directly from the water sample with nhexane. After drying and removing the bulk of the solvent, the insecticides are isolated from extraneous material by microcolumn adsorption chromatography. The insecticides are then analysed by gas chromatography. This method is a modification and extension of the procedures developed by Lamar et al. [30, 31]. For the analysis of insecticides in waters that are grossly polluted by organic compounds other than pesticides, a high-capacity clean-up procedure is used, as detailed in the Federal Water Pollution Control Administration Method for Chlorinated Hydrocarbon Pesticides in Water and Wastewater [32]. Mean recoveries obtained by this procedure for 12 insecticides from surface water samples range between 87.2% (Endrin) and 103% (Lindane). Lauren [22] extracted the sample containing 10–12g chlorinated insecticide with ethyl ether (50mL). The extract is concentrated to 1mL and placed on a column of deactivated silica gel (1g, 60–200 mesh) moistened with hexane (pesticide grade), and elution is effected with benzene–hexane (1:9) (6.5mL, followed by a further 6.5mL), the two fractions being collected separately. Each fraction is concentrated in a stream of air and injected into a gas chromatograph equipped with an electron-capture detector and a glass column (4ft (1.2m) × 3mm) packed with 3% of QF-1 and 2% of OV-1. Organophosphorus insecticides, which are retained on the silica, can be eluted as a third fraction with ethyl acetate–benzene (1:9). Croll [33] has given details of the use of back-flushing with electroncapture gas chromatography for the determination of organochlorine insecticides in water. Attention is paid to equalization of column resistances under operating and back-flushing conditions; baseline drift is thereby minimized. The system has been successfully used (with a variety of stationary phases, temperature ranging from 25 to 225°C and nitrogen flow rates from 25 to 200mL min-1). Kahanovitch and Lahav [9] used gas chromatography to study the occurrence of 12 organochlorine insecticides in water sources in Israel (α-BHC, γ-BHC, o,p’-DDD, p,p’-DDT, Dieldrin, Endrin, Heptachlor, Heptachlor epoxide, Thiodan, Treflan, Diazinon and Malathion). Water samples of 10L were extracted at the sampling sites with about 100mL hexane. After drying over anhydrous sodium sulphate, the extract was evaporated in a Buchi rotavapor apparatus; 2–3mL of hexane was then added to redissolve the insecticides. The clean-up procedure used was that described by Law and Goerlitz [34]. Effluent volumes of 0.5–2mL were collected and their amounts determined gravimetrically. The total
500
Insecticides and herbicides
effluent volume was usually 10mL. Pesticide mixtures of known concentrations were used to identify the eluent fraction in which each compound was eluted. The insecticide determination was made with a gas chromatograph equipped with an electron-capture detector. The glass columns used were of 4mm i.d. and 180cm length packed with (1) 5% OV-2% QF-1, and (2) 5% QF-1. Nitrogen was used as the carrier gas. The temperatures were: injection point, 210°C; column oven, 190°C; detector, 200°C. Concentrations were calculated from the peak height, since a linear relationship was found between the two. Due to the large water volumes used in the method, insecticide concentrations lower than 1ng L-1 could be detected without difficulty. Brodtmann [35] carried out a long-term study on the qualitative recovery efficiency of the carbon adsorption method versus that of a continuous liquid–liquid extraction method for several chlorinated insecticides. Comparative results obtained by electron-capture gas chromatography indicate that the latter method may be more efficient (Table 9.2). In this method river water samples were passed through carbon for a predetermined period. The carbon extract was then obtained by chloroform extraction of the carbon in a modified Soxhlet apparatus for 36h using glass distilled, pesticide grade petroleum ether. The neutral fraction of the chloroform extract was then prepared for gas chromatography by the methods of Breidenbach [36]. Brodtmann [35] used a continuous liquid–liquid extraction apparatus as described by Kahn and Wayman [37] and Goldberg et al. [38] for the extraction of nonpolar solutes from river water. Pesticide grade petroleum ether, used in all cases, was recycled internally (initial solvent charge of 350mL), Table 9.2 Overall recovery rates
Source: Reprinted from Brodtmann [35]. Copyright (1975) American Water Works Association.
Chlorinated insecticides
501
thereby continuously exposing essentially fresh solvent to the river water. A Florasil clean-up step using sequential elutions with 6% and 15% ethyl ether–petroleum ether solutions was employed. A dual-column gas chromatograph equipped with two tritium electron-capture detectors was employed by Brodtmann [35]. Both columns were acrylic glass (1.83m × 0.32cm i.d.). Column A was packed with 5% DC-260 on 80/100 DCMS Chromosorb W. Flow rate of carrier gas (5% methane–argon) through this column was 80mL min-1. Column B was packed with 1.5% OV-17–1.95% QF-1 on 80/100 Chromosorb W DCMS support. Flow rate of carrier gas (5% methane– argon) through this column was 50mL min-1. Both pairs of injectors, columns, and detectors were maintained at, respectively, 212, 184 and 204°C. Aspila et al. [39] reported the results of an interlaboratory quality control study involving five laboratories on the electron-capture gas chromatographic determination of 10 chlorinated insecticides in standards and spiked and unspiked sea water samples (Lindane, Heptachlor, Aldrin, γ-Chlordane, α-Chlordane, Dieldrin, Endrin, p,p’DDT, Methoxychlor and Mirex). The methods of analyses used by these workers were not discussed, although it is mentioned that the methods were quite similar to those described in the Water Quality Branch Analytical Methods Manual [40]. Both hexane and benzene were used for the initial extraction of the water samples. A summary of percentage recovery is given in Table 9.3. The standard deviation values are large because they contain all the errors, bias and deviations accumulated from the interlaboratory comparisons. Heptachlor revealed a poor recovery, which confirmed its degradation [41–45]. The degradation product [41–43] is known to be 1hydroxychlordene. Suzuki et al. [10] studied the determination of chlorinated insecticides in river and surface waters using high-resolution electroncapture gas chromatography with glass capillary columns. The compared resolution efficiencies of organochlorine insecticides and their related compounds with wall-coated open tubular (WCOT) and support-coated open tubular (SCOT) glass capillary columns with those of conventional packed glass columns. These columns were coated with silicone V-101 as the liquid phase. Applicabilities of the glass capillary column to environmental samples were investigated. An all-glass system was used to prevent thermal decomposition. The ‘resolution index’, i.e. peak height/half-width of peak of standard injected, generally increased in the following order: conventional packed glass column < WCOT glass capillary column ≤ SCOT glass capillary column. Excellent resolution of insecticides was obtained
502
Insecticides and herbicides
Table 9.3 Average percentage recovery of insecticides
Source: Reprinted with permission from Aspila et al. [39]. Copyright (1977) Association of Official Analytical Chemists. Notes a Mean recoveries exclude data for Mirex and Heptachlor. All data from laboratory 2 were excluded in calculating the average percentage recoveries. b s.d. is the standard deviation for the average percentage recovery (X): x is the mean value of 8 results. c No results were used because Heptachlor is too unstable.
with SCOT glass capillary columns and WCOT glass capillary columns. Log–log plots of the resolution index versus relative retention times compared to Aldrin were linear. These workers used a Shimadzu GC-5AIEE glass chromatograph equipped with a dual electron-capture detector (3H, 300mCi, foil type). Coiled SCOT glass capillary columns (23m × 0.28mm) were used. With SCOT glass capillary columns, the liquid phase was coated to the salt layer adhered to the inner wall of the glass capillary column. With WCOT glass capillary columns, the liquid phase was directly coated to the inner wall of the glass capillary column. Therefore, the surface area per unit of length was broader in SCOT glass capillary columns than in the WCOT glass capillary columns. The glass capillary column was connected to a holder. An OV-101 PGC (3% on Gas Chrom Q, 80–100 mesh, U-shaped, 2m × 3mm) was used [46]. The gas chromatographic
Chlorinated insecticides
503
conditions used were as follows: temperatures of column, injector and detector, for both conventional packed glass column chromatography and glass capillary column chromatography were 190, 210 and 200°C, respectively. For the packed column a flow rate of carrier gas (highly purified nitrogen gas, 99.9999+%) was 60mL min-1. The flow rates through the glass capillary column were both adjusted to 2mL min-1. The effluent from both glass capillary columns was scavenged to 60mL min-1 and entered into the electron-capture detector. The splitter ratio was 1:25 for SCOT glass capillary columns and 1:40 for WCOT glass capillary columns respectively. The levels of α-BHC, β-BHC, γ-BHC and δ-BHC in a 1L sample of river surface water were 0.002, 0.013, 0.006 and 0.016ppb, respectively, on the glass capillary column. The relative retention times of the individual insecticides and their related compounds compared to Aldrin, amounts injected and ‘resolution index’ of conventional packed glass columns and WCOT and SCOT glass capillary columns are shown in Tables 9.4 to 9.6, respectively. Among BHC isomers, greater enhancements in the ‘resolution index’ of γ-BHC were shown with WCOT glass capillary column and/or SCOT glass capillary column than with a conventional Table 9.4 Relative retention times (RRT) compared to Aldrin,a nanograms (ng) injected and ‘resolution index’ (RI) on OV-101 packed column
Source: Reprinted with permission from Suzuki et al. [10]. Copyright (1977) American Chemical Society. Note a Retention time of Aldrin was 4.19min.
504
Insecticides and herbicides Table 9.5 Relative retention times (RRT) compared to Aldrin,a nanograms (ng) injected and ‘resolution index’ (RI) on OV-101 WCOT glass capillary column
Source: Reprinted with permission from Suzuki et al. [10]. Copyright (1977) American Chemical Society. Note a Retention time of Aldrin was 6.25min.
glass packed column. Moreover, the ‘resolution index’ with a SCOT glass capillary column was about twice as much as that with WCOT glass capillary columns. Similar results were obtained in the case of aBHC and ß-BHC. Suzuki et al. [10] found that Endrin decomposes during hightemperature gas chromatographic analysis forming aldehyde and/or ketone derivatives. However, it appears that the Endrin does not decompose, or is not being adsorbed on the column wall of SCOT glass capillary columns. As for the results of DDT and its related compounds, no greater increases of the ‘resolution index’ in p,p’-DDE, p,p’-DDT and o,p’-DDT with a WCOT glass capillary column, in comparison with a conventional packed glass column, could be found. Significant enhancements of the ‘resolution index’ in p,p’-TDE and o,p’TDE with WCOT glass capillary columns were shown. Also, similar results were obtained with SCOT glass capillary column. Considerable improvements in the ‘resolution index’ of Mirex and Methoxychlor with a SCOT glass capillary column were shown in comparison with those of conventional packed glass columns and WCOT glass capillary columns. However, the peak of Photodieldrin, the photochemically
Chlorinated insecticides
505
Table 9.6 Relative retention times (RRT) compared to Aldrin,a nanograms (ng) injected and ‘resolution index’ (RI) on OV-101 SCOT glass capillary column
Source: Reprinted with permission from Suzuki et al. [10]. Copyright (1977) American Chemical Society. Notes Retention time of Aldrin was 4.99min. b No peak appeared. a
and microbially formed isomer of Dieldrin which was present in the field soil, could not be detected with a SCOT glass capillary column. This might be due to the thermal decomposition and/or adsorption of Photodieldrin to the support of SCOT glass capillary column. The ‘resolution index’ with WCOT glass capillary columns was smaller than with conventional glass packed columns. The resolution of insecticides and their related compounds on conventional glass packed columns, WCOT glass capillary columns, and SCOT glass capillary columns is shown in Figs 9.1 – 9.3. With conventional glass packed columns, broad peaks with greater retention times were shown, and the BHC isomer could not be separated. However, with WCOT glass capillary columns and SCOT glass capillary columns, good resolution of the BHC isomers was obtained. Aldrin, Heptachlor, Heptachlor epoxide and Mirex could not be separated on all the columns used. On the contrary, resolution of Dieldrin and p,p’-DDE could not be obtained. In general, better resolution was obtained with WCOT glass capillary columns.
506
Insecticides and herbicides
Fig. 9.1 Resolution of standard insecticides and their related compounds on conventional packed columns. Gas chromatographic conditions shown in text. I, a-BHC; 2, ß-BHC; 3, ?-BHC; 4, dBHC; 5, Heptachlor; 6, Heptachlor epoxide; 7, Aldrin; 8, Dieldrin; 9, Endrin; 10, p,p’-DDE; 11, p,p’-TDE; 12, p,p’-DDT; 13, o,p’-DDE; 14, o,p’-TDE; 15, o,p’-DDT; 16, Mirex; 17, Methoxychlor; 18, Photodieldrin. (Reprinted with permission from Suzuki et al. [10]. Copyright (1977) American Chemical Society.)
Fig. 9.2 Resolution of standard insecticides and their related compounds on WCOT glass capillary column. Peak numbers identical with those in Fig. 9.1. (Reprinted with permission from Suzuki et al. [10]. Copyright (1977) American Chemical Society.)
Sackmauereva et al. [14] have described a method for the determination of chlorinated insecticides (BHC isomers, DDE, DDT and hexachlorobenzene) in water. In this method the water sample (1–3L) is extracted with petroleum ether. A vacuum is used to transfer the water sample (separation funnel No. 1) and petroleum ether (separation funnel No. 2) into a glass spiral filled with 8mm glass balls. The petroleum ether layer is then concentrated to a
Chlorinated insecticides
507
Fig. 9.3 Resolution of standard insecticides and their related compounds on WCOT glass capillary column. Peak numbers identical with those in Fig. 9.1. (Reprinted with permission from Suzuki et al. [10]. Copyright (1977) American Chemical Society.)
volume of about 0.5mL using a vacuum, and purified on an alumina column (Woelm, neutral, activated by heating at 300°C for 3h and deactivated by adding 11% water). Thereafter, insecticides were eluted with 15% dichloromethane in petroleum ether. The eluate was concentrated in a vacuum rotary evaporator to a volume of 1mL and then used for gas–liquid chromatography [47–50], on a column operated under the following conditions: temperature of the column 180–200°C, temperature of the injection port 210°C, temperature of the electron-capture detector (63Ni) 200–225°C, nitrogen flow rate 60– 80mL min-1, EC detector voltage 20–70V. A column filled with 1.5% silicone OV-17 plus silicone oil (fluoralchylsiloxane) on Chromosorb W (80–100 mesh) is used for separation of the o,p’-DDT, p,p’-DDE, p,p’-DDD, and p,p’-DDT. a-BHC and hexachlorobenzene (HCB) have a common peak. They can be separated on a column filled with 2.5% silicone oil XE-60 (ß-cyanoethylmethylsilicone) on Chromosorb W (80–100 mesh). Using the gas chromatography methods, Sackmauereva et al. [14] obtained from spiked samples the four BHC isomers at 93–103.5% recovery. Both DDT and DDE were yielded in 85.6–94% from water. Surface water from the Danube was examined for the content of BHC, DDE and DDT residues. The average content of ?-BHC was 0.117µg L-1, and that of ß-BHC 0.040µg L-1, and that of the other BHC isomers plus HCB was 0.049µg L-1. The average content of DDE was 0.050µg L-1 and that of DDT 0.125µg L-1. Thompson et al. [18] have described a gas chromatographic procedure for the multiclass, multiresidue analyses of organochlorine insecticides in water. It involves extraction with methylene chloride, separation into
508
Insecticides and herbicides
groups on a partially deactivated silica gel column, and sequential elution with different solvents. Final determinations of halogenated compounds and derivatized carbamates are made by gas chromatography with electron-capture detection, and for organophosphorus compounds a flame photometric detector is used. This study included 42 organochlorine insecticides, 33 organophosphorus insecticides and seven carbamate herbicides. These are discussed here as such mixtures are likely to be encountered in actual samples. The work on organophosphorus and carbamate herbicide types are referred to again in the appropriate chapters. Thompson et al. [18] point out that only a few methods have been published which are ideally suited to the estimation of a wide variety of pesticidal compounds such as organochlorine and organophosphorus insecticides and herbicides. Table 9.7 is presented to illustrate the dearth of broadly applicable multiresidue methods in the literature. Of these references, only five are intended as multiclass, multiresidue procedures, outstanding amongst which is the work of Sherma and Shafik [51]. To correct this deficiency Thompson et al. [18] developed the multiclass, multiresidue method which will provide the analyst with a means of simultaneously monitoring a water sample for a wide variety of pesticides. The concentrated methylene chloride extract of the water sample is applied to a silica gel column, which is then eluted with n-hexane to provide fraction I, then with 60% v/v benzene–40% v/v n-hexane to provide fraction II, then with 5% v/v acetonitrile–95% v/v benzene to provide fraction III and, finally, with 25% v/v acetone–75% v/v methylene chloride to provide fraction IV. The fourth fraction is only necessary if there is reason to suspect the presence of Crufomate, Dimethoate, Mevinphos, Phosphamidon or the oxygen analogues of Diazinon or Malathion. The majority of the organochlorine insecticides will be detected in fractions I and II with a few of the more polar compounds in fraction III (Table 9.8). Most of the organophosphorus insecticides will be found in fractions I and II, a very few in fraction IV and none in fraction I. Carbamate insecticides are found in fractions II and III. Of the organochlorine insecticides examined a high proportion of reproducible recoveries of 80% or better were obtained in the majority of cases. Thompson et al. [18] used a gas chromatograph with electroncapture and flame photometric detectors, the latter operated in the phosphorus mode at 526nm. Columns were 1.8m × 4mm i.d., borosilicate glass packed with 1.5% OV-17–1.95% OV-210 or 5% OV-210, both coated on Gas-Chrom Q (80–100 mesh). Other workers [5] have used electron-capture gas chromatography coupled with the use of glass capillary columns for the separation of chlorinated insecticides. Figure 9.4 shows separations achieved of 15
Source: Reprinted with permission from Thompson et al. [18]. Copyright (1977). Springer-Verlag, New York.
Table 9.7 Multiresidue methods
510
Insecticides and herbicides
Table 9.8 Recoveries of 20 organochlorine compounds
Source: Reprinted with permission from Thompson et al. [18]. Copyright (1977) Springer Verlag, New York. Note a Non-linear response.
chlorinated insecticides on a WCOT glass capillary column containing polyphenylmethyl silicone gum (CPtm Sil7). 9.1.1.2 Gas chromatography–mass spectrometry Gas chromatography–mass spectrometry involving chemical ionization mass spectrometry has been used to detect and confirm the presence of Kepone (chlorodecone), Kepone photoproducts and a conversion product of Kepone in environmental samples [58]. 9.1.1.3 Thin-layer chromatography Thin-layer chromatography has been used by several workers to confirm the identity of chlorinated pesticides [57, 59–62, 64, 110]. Arias et al. [63] have described a method for the determination of organochlorine
Chlorinated insecticides
511
Fig. 9.4 Chromatogram of 15 chlorinated insecticides (test mixture 50 × diluted) (Copyright (1979) Chromopak International BV [5].)
insecticide residues in river water. The method involves extraction, Florasil column clean-up, and analysis of the extract by thin-layer chromatography on silica gel G or alumina with hexane or hexane– acetone (49:1) as solvent, or gas chromatography on a polar column of 10% DC-200 on Chromosorb W HMDS and on a semipolar column of 5% of DC-200 plus 7.5% of QF-1 on Chromosorb W, with electron-capture detection. Bevenue et al. [65] point out that when a comparatively small sample of water (1 gallon (4.5L) or less) is extracted for detection of pesticides and submitted to thin-layer chromatography, the use of spray reagents may not be practical. The spots are therefore extracted, and the concentrated extracts are submitted to gas chromatography. Extraneous interferences are magnified on the recorder chart unless special precautions are taken to exclude impurities from the organic solvents or from the apparatus used. The necessary pretreatment of glassware and silica gel is discussed and described. Plastic ware and filter paper should not be used. Suzuki et al. [66, 67] separated chlorinated insecticides into two groups by column chromatography and then into three further groups by thin-layer chromatography. Individual insecticides were then isolated
512
Insecticides and herbicides
and determined by gas chromatography. The members of the two groups separated by column chromatography were: first-division insecticides – Aldrin, DMC ethylene (1,1-bis-(4-chlorophenyl)ethylene), chlorfensulphide, DDS (bis-(4-chlorophenyl)disulphide), o,p’- and p,p’DDT, Heptachlor, Quintozene, Isobenzan and Tetrasul; second-division insecticides – alpha-, beta- and gamma-BHC, Triflurin, Nitrofen, Dieldrin and Dicofel. Sackmauereva et al. [14] used thin-layer chromatography on silica plates to confirm the identity of chlorinated insecticides previously identified by gas chromatography. The compounds can be separated by single or repeated one-dimensional development in n-heptane or in n-heptane containing 0.3% ethanol. The plate was dried at 65°C for 10min and detected by spraying with a solution of silver nitrate plus 2-phenoxyethanol. Thereafter, the plate was dried at 65°C for 10min and illuminated with an ultaviolet light (λ. = 254nm) until spots representing the smallest amounts of standards were visible (10–15min). The pesticide residues may be evaluated semiquantitatively by simple visual evaluation of the size and of the intensity of spot coloration, and by comparing extracts with standard solutions. Armstrong and Terrill [68] used aqueous solutions of sodium sulphate with polyamide to separate p,o’-DDT, p,p’-DDD and p,p’-DDE and decachlorobiphenyl on alumina-coated thin-layer sheets. 9.1.1.4 High-performance liquid chromatography Willmott and Dolphin [69] applied their high-performance liquid chromatography–electron-capture detector approach to the determination of chlorinated insecticides in surface waters. Because the concentration of pesticides in most surface waters is less than 1(µg L-1, some form of extraction and concentration from large volumes of water is necessary before analysis is possible. These workers applied a conventionally coated chromatographic support as a reverse liquid– liquid partitioning filter to the extraction of pesticides. They used uncoated polyether polyurethane foam; 0.5g of the flexible foam were inserted into a 10mm i.d. quartz tube and cleaned by washing with consecutive 100mL aliquots of acteone, n-hexane, ethanol and distilled water. 9.1.1.5 Nuclear magnetic resonance spectroscopy Keith and Alford [70] have reviewed the applications of NMR spectroscopy to the determination of organochlorine pesticides (57 references).
Chlorinated insecticides
513
9.1.1.6 Yeast hexokinase assay Sadar and Guilbauelt [71] have described an assay method in which down to 1µmol of chlorinated insecticides, organophosphorus insecticides, and herbicides and fungicides can be determined with an error of about 5%. 9.1.1.7 Photodegradation technique Chau [72] has investigated photodegradation products of Endrin including hexachloroketone and pentachloroketone. The residues obtained after clean-up on a Florasil column were heated with potassium t-butoxide in t-butyl alcohol. A procedure involving ultraviolet irradiation followed by gas–liquid chromatographic detection of photodecomposition products has been used by Erney [73] to confirm the identity of selected organochlorine insecticides. Erney [73] studied the photodecomposition product patterns of 33 chlorinated insecticides. The attractiveness of this technique lies in the fact that it offers a means of independent identification of insecticides. The degree of photochemical reaction is dependent on wavelength intensity and time of irradiation as well as the physical state, which must be controlled to give consistent reactions suitable for identification. 9.1.1.8 Miscellaneous Novikova [74] has reviewed the literature (209 references) covering the extraction, clean-up, and analyses of organochlorine and organophosphorus insecticides in water. Laubereau [75] has discussed the requirements for a standard method for determination of high molecular weight organochlorine and related compounds in water and the contents of the final draft prepared by a working party on this subject is now being reviewed. This method has been evaluated by round-robin test among German laboratories. The US Environmental Protection Agency [76] has also published methods for organochlorine pesticides in water and waste water. This is the official US approved method for the determination of organochlorine pesticides in an aqueous medium. It includes advice on the collection of the sample, choice of correct glassware, preparation of stock standards, solvents and reagents, concentration of samples, preparation of gas chromatographic columns, and injection devices, and gives details of the gas chromatographic analytical method. The removal of sulphur from water samples prior to the gas chromatographic determination of chlorinated insecticides has been
514
Insecticides and herbicides
discussed [77, 78]. Woodham and Collier [77] used copper, whilst Baird et al. [78] used a column packed with a mixture of 4% of OV-17, 47% of QF-1, and 1% of DC-200 on Gas-Chrom G. An all-glass apparatus for sampling water samples for chlorinated insecticide analyses has been described [79]. Interference effects on the gas chromatogram have been reported when water samples are filtered through ordinary filter paper clamped in a PTFE gasket [80]. Levi and Nowicki [81] have overcome this interference by using a filtration apparatus constructed in stainless steel with a fritted borosilicate glass filtration disc. An automatic sampler has been used [82] to observe an Endosulphan wave in the river Rhine. The automatic sampler at the gauging station was programmed to provide 2h mixed samples composed of 125mL portions taken every 10min. The resulting samples, together with the 24h mixed samples for the preceding and succeeding periods were analysed for Endosulphan by gas–liquid chromatography. The results show a distinct peak in the Endosulphan concentration with a maximum recorded value of 276mg L-1 or about 100 times the normal background level. Estimation of the total mass flow with the aid of flow measurements indicated that 40–50kg Endosulphan passed Koblenz during August, compared with a baseline value of only 5kg. Le Roy and Goerlitz [83] have described the use of microcolumns (4cm × 5mm) with deactivated alumina, silica gel and Florasil for the clean-up of chlorinated insecticide prior to gas chromatography. Weil and Quentin [84] used silica gel for the clean-up of water samples prior to analysis for organochlorine insecticides. Balayanis [85] used surface modified adsorbents in a clean-up technique for analysis of DDT and γ-BHC in water prior to gas chromatography. The adsorbents comprised silica gel modified by treatment with trichorooctadecylsilane or with dichlorodimethylsilane. These two products effected substantial clean-up, but also displayed different selectivities towards pesticides. p,p’-DDE was strongly retained by the first product, and required a second elution with 10mL of hexane to recover it all, whereas with the second product, almost the whole of the added p,p’-DDE was recovered in the first 10mL of hexane. p,p’-DDT was strongly retained by both products, as was ?-BHC. With 10mL of hexane-acetone (19:1), ?-BHC could be eluted readily, without interfering peaks, from the first product. 9.1.1.9 Preconcentration Preconcentration methods used in the determination of organochlorine insecticides in natural waters are reviewed in Table 9.9. Many of these
Chlorinated insecticides
515
Table 9.9 Preconcentration methods for organochlorine pesticides prior to gas chromatography
methods enable organochlorine insecticides to be determined at the submicrogram per litre level. 9.1.1.10 Individual organochlorine insecticides Dichlorvos Shevchuk et al. [112] investigated extraction efficiency by extractive chromatography of the extraction of Dichlorvos using hexane, and
516
Insecticides and herbicides
hexane with benzene and toluene. Hexane alone was not effective, the degree of extraction at pH2.5–7.5 reached a maximum 26%. The effect of sodium, calcium, aluminium, lanthanum, gallium, yttrium and gadolinium and nitrate on the extraction with hexane showed that salting out significantly affected the distribution of Dichlorvos between the aqueous and the organic phase. Mixtures of solvents were more effective at extracting the Dichlorvos, and a synergistic effect was observed. The most effective mixture was hexane–benzene at a volumetric ratio of 2:3. Addition of 3.0M aluminium nitrate increased the extraction efficiency to 91%. Surface active agents and organic anions decreased the salting-out effect. Gas chromatography with flame ionization detector was used to determine the concentration of Dichlorvos and compared with a thin layer chromatographic method. Preconcentration allowed an increase in the sensitivity of the determination by an order of magnitude. Hexochlorocyclohexone and hexachlorobenzene isomers Malaiyandi et al. [28] have discussed some of the difficulties associated with the gas chromatographic determination of ultra trace levels of pesticide residues in environmental samples. A supposedly operator-independent method was tested for the recovery of hexachlorocyclohexane isomers from water. This recovery method, designed to reduce losses during extraction and evaporation, was tested on samples containing 30–350ng L -1 of alpha-, beta-, gamma- and delta-hexachlorohexane isomers employing a Kuderna–Danish evaporator and modified Snyder column. Mean recoveries of alpha-, beta- and gammahexachlorocyclohexane isomer were better than 90%. However, recovery of the delta-hexachlorocyclohexane isomer was 63–85%. This was attributed to degradation at the injection port (250°C). Significant differences in operator variability were observed. DDT Taylor et al. [11, 113] isolated DDT from water and determined it by gas chromatography. This method is based on the extraction of pesticides from water with light petroleum, purification of the concentrated extract on a Florasil column followed by gas chromatography at 195°C on a column (160cm × 3mm) packed with 5% of DC-11 on Chromosorb W AW DMCS, with argon (45mL min-1) as the carrier gas and tritium electroncapture detector. For water containing appreciable amounts of suspended solids, the sample was filtered, and the solids were collected on a filter, dried, and extracted in a Soxhlet apparatus for 5h with acetone.
Chlorinated insecticides
517
Deubert [114] has discussed the sources of compounds which interfere in the analyses in water for DDT and Dieldrin by gas electroncapture chromatography. Nitration of these insecticides eliminated their peaks so that background interference peaks could be studied. Trichlorphon Trichlorphon has been determined [115] in water, by solvent extraction followed by gas chromatography on a glass column (6ft × 0.375in (1.8m × 9.5mm)) packed with 16% of XF-1150 on Chromosorb W-AW operated at 125°C, with a carrier gas flow of 60mL min-1 and a flame photometric detector operated in the phosphorus mode. Average recoveries were 96%, and down to 0.002ppm of Trichlorphon could be determined in water. Fuka et al. [116] have studied the decomposition and determination of Trichlorphon and Dichlorvos in pond water. Gas chromatographic evidence indicates that Trichlorphon undergoes partial conversion to Dichlorvos in a few days. Aldrin Lester and Smiley [117] developed a rapid method for identifying Aldrin in the presence of sulphur by electron-capture gas chromatography. The presence of elementary sulphur in hexane extracts of water or sediment produces a response during gas chromatography with electron detection that is similar to the response produced by Aldrin. It has been established that sulphur and Aldrin can be separated on a glass column (6ft × 0.25in. o.d. (1.8m × 6mm)) packed with 3% OV-17 on Chromosorb W-HP. The column is operated at 180°C and the carrier gas flow is 90mL min-1. Hexachlorophane In a method [118] for determining this insecticide, the hexachlorophane is converted into its p-anisate ester, which is subjected to chromatography on a column (2ft (0.6m) × 2.3mm) packed with Sil-X silica (36– 40µm) with 1-chlorobutane as solvent and detection by UV absorption. Calibrations are made over the range 40–280ng of the derivative. The coefficient of variation at the 20ng level is 1.0%. Endosulphan Chau and Terry [2] in a series of papers have discussed gas chromatographic methods for the determination of a- and ßEndosulphan. The procedure involved heating the sample on a 2cm column of alumina impregnated with concentrated sulphuric acid and
518
Insecticides and herbicides
heated 1.5h at 100°C to convert Endosulphan to an ether which was examined by gas chromatography. Down to 0.003ng of the insecticide could be detected per millilitre of water extract. The method was subsequently applied to the determination of 10pg of Heptachlor in 1L of water [119]. An automatic sampler has been used [82] to observe an Endosulphan wave in the river Rhine. The automatic sampler at the gauging station was programmed to proved 2h mixed samples composed of 125mL portions taken every 10min. The resulting samples, together with the 24h mixed samples for the preceding and succeeding periods were analysed for Endosulphan by gas–liquid chromatography. The results show a distinct peak in the Endosulphan concentration with a maximum recorded value of 276ng L-1 or about 100 times the normal background level. Estimation of the total mass flow with the aid of flow measurements indicated that 40–50kg Endosulphan passed Koblenz during August, compared with a baseline value of only 5kg. Dieldrin and Endrin Woodham et al. [120] converted Dieldrin and Endrin to chemical derivatives prior to gas chromatographic determination of these substances in water. An aliquot of extract, after appropriate clean-up and evaporation to dryness, is treated with 1mL of conversion reagent (10% solution of boron trichloride in 2-chloroethanol) in a centrifuge tube, which is then placed, unstoppered in a water bath at 90°C and left for 3h or for 10min for a sample that may contain Dieldrin, or Endrin, respectively, or for 2h for a mixture. The tube is cooled, 5mL of hexane and 10mL of 7% aqueous sodium sulphate are added, and the contents are mixed and left for the phases to separate. The hexane phase is analysed on a column of OV-17–QF-1 on Gas Chrom Q with electroncapture detection (tritium source). Down to 0.01ppm of either pesticides (0.01 part per 109 in water) could be detected. Methoxychlor Fredeen et al. [121] have described a gas chromatographic method for determining Methoxychlor residues at sub-ppm level in river water following injections of this substance into a river. Two litre aliquots of water samples were filtered under suction to remove suspended soils. The filtrate was extracted with 200mL of hexane three times. The combined hexane extract was dried over anhydrous sodium sulphate, concentrated to a suitable volume and analysed by electron-capture– gas– liquid chromatography.
Chlorinated insecticides
519
Retention time for Aldrin was 3.0min. Retention time of other organochlorine insecticides relative to Aldrin were: Heptachlor 0.94; Heptachlor epoxide 1.81; p,p’-DDE 2.47; Dieldrin 2.81; p,p’-DDD 3.25; p,p’-DDT 3.78; and p,p’-Methoxychlor 7.25. A combination of gas chromatography with mass spectrometry was used to confirm the identity of the Methoxychlor peak on the gas chromatogram. Toxaphene (Camphechlor) Hempel et al. [122] have described a colorimetric method for determining down to 15 parts per 109 of this pesticide in water. Toxaphene is first extracted into hexane then the extract washed with concentrated sulphuric acid then chromic acid then water. Hexane is removed in vacuo and the residue taken up in acetone prior to evaporating to dryness in vacuo. Pyridine is then added and the mixture heated for 30min at 100°C, and aqueous ethanol and sodium hydroxide are added. The extinction of the coloured reaction product is measured at 550nm. Hughes and Lee [123] have used gas chromatography to determine Toxaphene in water and fish samples. 9.1.2 A queous precipitation 9.1.2.1 Gas chromatography Engst and Knoll [124] used hexane extraction gas chromatography to study the occurrence in rain water of down to 0.001µg L-1 of p,p’-DDT, p,p’-DDE and p,p’-TDE. Samples (500mL) were extracted by shaking with hexane (100, 50 and 50mL) for 1min each time. The combined hexane phases were dried with sodium sulphate and evaporated to between 1 and 5mL in a rotary evaporator. The extract (1mL) was injected, without further clean-up, on to a gas chromatographic assembly consisting of one column (6ft × 0.25in (1.8m × 6mm)) packed with 5% QF-1 on Varaport 30 (100–120 mesh) and a second column (3ft × 0.25in (0.9m × 6mm)) packed with 4% of OV-17 on AW-DMCS Chromosorb W (60–80 mesh) both operated at 180°C with nitrogen as the carrier gas (30mL min-1) and a tritium electron-capture detector. 9.1.3 Sea water 9.1.3.1 Gas chromatography Earlier work by Wilson et al. [125, 127] has indicated that organochlorine pesticides were not stable in sea water, as indicated in Table 9.10.
520
Insecticides and herbicides
Table 9.10 Stability of pesticides in natural sea water (salinity 29.8ppt)
Notes a Metabolites of parent compound. b From the seventeenth day onward, two unidentified peaks appeared on the chromatographic charts after Aldrin had eluted.
Since petroleum ether was the solvent used in these earlier studies for extracting DDT from sea water, Wilson and Forester [126] initiated further studies to evaluate the extraction efficiencies of other solvent systems, viz. petroleum ether, 15% ethyl ether in hexane followed by hexane, or methylene chloride. Wilson and Forester [126] discussed the determination of Aldrin, Chlordane, Dieldrin, Endrin, Lindane, o,p and p,p’ isomers of DDT and its metabolites, Mirex and Toxaphene in sea water. The concentrated solvent extracts were analysed by electron-capture gas chromatography using columns of different liquid phases. The following columns were used: DC-200, QF-1, EGS, OV-101, mixed DC-200–QF-1 and mixed OV101–OV-17. Just prior to extraction, all samples were fortified with o,p’DDE to evaluate the integrity of the analysis. The recovery rates of o,p’DDE in all tests were greater than 89%, indicating no significant loss during analyses. After initial sampling, the bottles were sealed and incubated at 20°C under controlled light conditions. Dark samples were extracted at various time intervals. At time zero a recovery of 93% p,p’-DDT was obtained from sea water (salinity 16–21ppt) regardless of the solvent used. After 6 days exposure in daylight the recovery decreased to 67% in the case of petroleum ether or 15% with diethyl ether in n-hexane and 76% in the case of methylenechloride. The results in Table 9.11 show the effect of exposure time, light and temperature conditions of the recovery of p,p’-DDT from sea water. Recoveries of p,p’-DDT from distilled water under the same conditions were in the range 90–92% with all the extraction solvents studied. Since distilled water is devoid of particulate matter, this study suggests that DDT may be absorbed or adsorbed on to plankton or
Chlorinated insecticides
521
Table 9.11 Percentage recovery of p,p’-DDT from sea water incubated under different light and temperature conditions
particulate matter in sea water and the sorbed material was not removed resulting in low recoveries of DDT. This would explain the initially high extraction efficiency of DDT followed by the decline in recovery as DDT was associated with the particulate phase. Since methylene chloride was the most polar solvent used, it would have a greater affinity for removing sorbed DDT. These experiments support the work of other investigators in that DDT and other insecticides are extremely hydrophobic and can easily be absorbed or adsorbed by suspended matter from liquid solutions. Results obtained using petroleum ether as extraction solvent support the concept that physical or chemical transformations of insecticides altered the extraction efficiencies of the solvent and prevented complete recovery of the compounds. It is difficult to relate laboratory findings directly to those of the estuary or open oceans. However, the laboratory data illustrate clearly some problems that could be encountered in monitoring sea water samples for insecticide pollution. The conventional analyses of water samples by liquid–liquid extraction techniques may provide invalid data if suspended matter is not considered. Standardized methods are needed to analyse the water column and suspended material separately. Wilson [127] showed that liquid–liquid extraction of estuarine water immediately after addition of DDT gave acceptable recovery levels with all solvent systems tested, but analyses carried out several days later gave only partial recovery owing to adsorption of DDT on suspended matter. 9.1.3.2 Miscellaneous Liquid scintillation counting of [14C]DDT has been used to study the pickup and metabolism of DDT by fresh water algae and to determine DDT in sea water [127]. Picer et al. [128] described a method for measuring the radioactivity of labelled DDT contaminated sediments in relatively large volumes of
522
Insecticides and herbicides
water using a liquid scintillation spectrometer. Various marine sediments, limestone and quartz in sea water were investigated. External standard ratios and counting efficiencies of the systems investigated were obtained as was the relation of efficiency factor to external stan dard ratio for each system studied. 9.1.3.3 Preconcentration Harvey [129] has examined ion-exchange resins (Amberlite XAD-2) as an absorption medium for chlorinated insecticides in sea water. 9.1.4 Potable wates 9.1.4.1 Gas chromatography Sackmauereva et al. [14] have described the method given below, for the determination of chlorinated insecticides (BHC isomers, DDE, DDT and hexachlorobenzene) in potable waters. The water sample (1–3L) is extracted with three portions of petroleum ether (boiling point 30–40°C). The petroleum ether layer is then concentrated to a volume of about 0.5mL using a vacuum and purified on an alumina column (Woelm, neutral, activated by heating at 300°C for 3h and deactivated by adding 11% water). Thereafter, insecticides were eluted with 15% dichloromethane in petroleum ether. The eluate was concentrated in a vacuum rotary evaporator to a volume of 1mL and then used for gas–liquid chromatography. When the individual insecticides are present in the solution in such a concentration range, the electron capture responds nearly uniformly to all insecticides. A column filled with 1.5% silicone OV-17 plus silicone oil (fluoralkylsiloxane) on Chromosorb W (80–100 mesh) is used for separation of the BHC, alpha, beta, gamma and delta isomers (hexachlorocyclohexane), o,p’-DDT, p,p’DDE, p,p’-DDD and p,p’-DDT. a-BHC and hexachlorobenzene (HCB) have a common peak. They can be separated on a column filled with 2.5% silicone oil XE-60 (ß-cyanoethylmethylsilicone) on Chromosorb W (18–800 mesh). Using the gas chromatography methods, Sackmauereva et al. [14] obtained from spiked sample the four BHC isomers at 93–103.5% recovery. Recoveries of DDT and DDE were 85.6–94% from water. Purification on a Florasil column was used in determining chlorinated insecticides unstable at low pH (Aldrin, Dieldrin). The type and activity of Florasil influence the yield and accuracy of the method. Therefore, the activity of the adsorbent had to be verified and adjusted. From the results of the analyses of 92 potable water samples the average content
Chlorinated insecticides
523
of γ-BHC was 0.069µg L-1, that of ß-BHC was 0.023µg L-1 and that of the other isomers (α + δ) of BHC + HCB (hexachlorobenzene) 0.018µg L-1. The average content of DDE in potable water was 0.022µg L-1 and that of DDT 0.042µg L-1. The Department of the Environment (UK) [130] has used a tentative method for the determination of organochloride insecticides and PCBs in potable and natural waters. The first part of this method is concerned with the extraction and determination of the amounts of individual substances present; the second part is concerned with methods for verifying the identity of the various substances quantified in the first part. Representative data are presented for a number of typical insecticides using different chromatographic stationary phases. Ronsen [131] has discussed a systematic analytical scheme for the determination of organic pollutants including Toxaphene insecticide and PCBs in potable water. The pollutants are divided into volatile and extractable compounds, and the extraction further divided into acid and base extractables and into those capable of flame ionization or electroncapture detection. The pesticides, Toxaphenes and Arochlors are combined into one category. 9.1.4.2 Thin-layer chromatography Mosinska [132] has described a semiquantitative thin-layer chromatographic method for the determination of Trichlorphon in potable water. The sample is extracted with redistilled chloroform. The extract is dried with sodium sulphate, reduced in volume to 5mL in vacuo and then evaporated to dryness in a stream of air. The residue is dissolved in acetone and chromatographed on chloride-free silica gel G plates (activated at 100°C for 1h) with benzene–methanol (17:3) as solvent. The spots, revealed with ammoniacal silver nitrate in acetone, are compared with those of standards for semiquantitative determination. The detection limit is 0.02mg L-1 and the efficiency of extraction is 70%. Sackmauereva et al. [14] used thin-layer chromatography on silica plates to confirm the identity of chlorinated insecticides previously identified by gas chromatography. The compounds can be separated by single or repeated one-dimensional development in n-heptane or in nheptane containing 0.3% ethanol. The plate was dried at 65°C for 10min and detected by spraying with a solution of silver nitrate plus 2phenoxyethanol. Thereafter, the plate was dried at 65°C for 10min and illuminated with an ultraviolet light (λ = 254nm) until spots representing the smallest amounts of standards were visible (10–15min). The pesticide residues may be evaluated semiquantitatively by simple visual evaluation of the size and of the intensity of spot coloration and by comparing extracts with standard solutions.
524
Insecticides and herbicides
Sherma and Slobodien [133] determined Chloropyrifos insecticide and its metabolite 3,5,6-trichloro-2-2-pyridinol in potable water at 5µg L1 by thin-layer chromatography. Preadsorbent silica gel layers were used for resolution and silver nitrate for detection prior to reflectance scanning. Recovery of 3,5,6-trichloro-2,2-pyridinol from water was 84%. Recovery of Chloropyrifos from potable water was 87.5%. 9.1.5 Sewage effluents 9.1.5.1 Gas chromatography Laseter et al. [134] and Kaiser [135] have both utilized gas chromatography and mass spectrometry for the determination of Mirex. Kaiser [135] has pointed out that under standard gas chromatographic conditions, the Mirex peak is superimposed on that of the PCBs and, as a result, the presence of Mirex may have been interpreted by several workers as a PCB isomer. He used a computer-controlled gas chromatographic–mass spectrometric system to positively identify Mirex and distinguish it from other highly chlorinated insecticides that could have been present in the samples including Aldrin, Chlordane, Dieldrin, Endrin, Endosulphan, Heptachlor, Kepone (chlordecone), Pentac and Toxaphene. 9.1.5.2 Miscellaneous Andrade and Wheeler [136] have studied the biodegradation of Mirex (dechlorane C H ) by sewage sludge or organisms utilizing 14C-labelled 10 12 Mirex. They did not succeed in identifying the metabolites. 9.2 Mixtures of chlorinated insecticides and polychlorobiphenyls 9.2.1 Natural waters 9.2.1.1 Gas chromatography The previous pages dealt with methods for chlorinated insecticides alone. In actual practice, environmental samples which are contaminated with PCBs are also highly likely to be contaminated with chlorinated insecticides. Many reports have appeared discussing co-interference effects of chlorinated insecticides in the determination of PCBs and vice versa and much of the more recent published work takes account of this
Mixtures of chlorinated insecticides and polychlorobiphenyls
525
fact by dealing with the analysis of both types of compounds. This work is discussed below. PCBs have gas chromatographic retention times similar to the organochlorine insecticides and therefore complicate the analysis when both are present in a sample. Several techniques have been described for the separation of PCBs from organochlorine insecticides. A review of these methods has been presented by Zitko and Choi [137]. These techniques are time-consuming and, in general, semiquantitative. In addition, differential adsorption or metabolism of the Arochlor isomers in marine biota prevent accurate analysis of the PCBs. The gas chromatographic determination of chlorinated insecticides together with PCBs is difficult. Chlorinated insecticides and PCBs are extracted together in routine residue analysis, and the gas chromatographic retention times of several PCB peaks are almost identical with those of a number of peaks of chlorinated insecticides, notably of the DDT group. The PCB interference may vary, because the PCB mixtures used have different chlorine contents, but it is common for PCBs to be very similar to many chlorinated insecticides and the complete separation of chlorinated insecticides from PCBs is not possible by gas chromatography alone [138–142]. Figure 9.5 illustrates the possibility of the interference of DDT-type compounds in the presence of PCBs. In an early paper on the determination of PCBs in water samples which also contain chlorinated insecticides, Ahling and Jensen [143] pass the sample through a filter containing a mixture of Carbowax
Fig. 9.5 Interference of DDT-type compounds in the presence of PCBs. Glass capillary column: 5% QF-1 on Gas Chrom Q, 100–120 mesh. Solid line: 1 = p,p’-DDT; 2 = p,p’-DDD; 3 = o,p’-DDT; 4 = o,p-DDD; 5 = p,p’-DDE; 6 = p,p’-DDMU; 7 = o,p’-DDE; 8 = o,p-DDMU. Broken line: PCB Chlophen A 50. (Reprinted with permission from Zitko and Choi [137]. Copyright (1971) Fisheries Research Board, Canada.)
526
Insecticides and herbicides
4000 monostearate on Chromosorb W. The adsorbed insecticides are eluted with light petroleum and then determined by gas chromatography on a glass column (160cm × 0.2cm) containing either 4% SF-96 or 8% QF-1 on Chromosorb W pretreated with hexamethyldisilazane, with nitrogen as carrier gas (30mL min-1) and a column temperature of 190°C. When an electron-capture detector was used the sensitivity was 10ng of Lindane per cubic metre, with a sample size of 200L. The recoveries of added insecticides range from 50 to 100%; for DDT the recovery is 80% and for PCBs 93–100%. Dolan and Hall [144] have described a Coulson electrolytic conductivity detector of enhanced sensitivity for the gas chromatographic determination of chlorinated insecticides in the presence of PCBs. The detector was modified by the replacement of the silicone-rubber septum and stainless steel fitting at the exit of the pyrolysis furnace with PTFE fitting, by the reduction in diameter of the PTFE transfer tube, and by the replacement of the 4mm (i.d.) reaction tube with one of 0.5mm. These modifications reduced hydrogen chloride adsorption and tailing, and improved sensitivity and reproducibility; sensitivity was also enhanced by increasing the cell voltage to 44V. Musty and Nickless [145] used Amberlite XAD-4 for the extraction and recovery of chlorinated insecticides and PCBs from water. In this method a glass column (20cm × 1cm) was packed with 2g of XAD-4 (60–85 mesh) and 1L of tap water (containing one part per 109 of insecticides) was passed through the column at 8mL min-1. The column was dried by drawing a stream of air through, then the insecticides were eluted with 100mL of ethyl ether–hexane (1:9). The eluate was concentrated to 5mL and was subjected to gas chromatography on a glass column (5.5ft (1.7m) × 4mm) packed with 1.5% OV-17 and 1.95% QF-1 on Gas-Chrom Q (100–200 mesh). The column was operated at 200°C, with argon (10mL min -1 ) as carrier gas and a 63Ni electron-capture detector (pulse mode). Recoveries of BHC isomers were 106–114%; of Aldrin, 61%; of DDT isomers, 102–144%; and of PCBs, 76%. Elder [146] evaluated mixtures of PCBs and DDE in terms of mixtures of commercial preparations from peak heights of packed column gas chromatograms using a programmable calculator. They proposed a method for evaluating gas chromatograms of multicomponent PCB mixtures and superimposed single components simultaneously. Apparent concentrations relative to calibration mixtures are assigned to a number of suitable peaks, and the apparent concentrations are related to the true concentrations by a set of linear equations which are solved by least squares approximation.
Mixtures of chlorinated insecticides and polychlorobiphenyls
527
Södergren [147] investigated the simultaneous detection of PCBs, chlorinated insecticides, and other compounds by electron-capture and flame ionization detectors combined in series using an open tube capillary column. Södergren [147] points out that much of the earlier work on combining detectors has utilized splitting the sample before a parallel detection system. As a result the sensitivity of the system is decreased, since only part of the injected sample is fed to each detector. Furthermore, it may be difficult to quantify the detected compounds since the splitting ratio, usually estimated by measurement of the carrier gas stream, may be affected by the size of the molecules eluted from the column. Södergren [147] combined the electron-capture detector and the flame ionization detection in series to obtain a dual detection system capable of simultaneous detection of environmental pollutants of different character, e.g. organochlorine residues and oil and lipid constituents in samples from aquatic environments. In order that the limit of detection should not be adversely affected when using capillary columns, a splitless system without a scavenging gas was used. Södergren [147, 148] connected a modified all-glass electroncapture detector to the flame ionization detector of a gas chromatograph. Since electron-capture is a non-destructive process, the effluent from the column passes undisturbed through the electron capture detector. The effluent was then directed to the jet-tip of the flame ionization detector by means of a glass capillary tube. The detector system was connected to the column by means of a PTFE tube. Glass columns (190cm × 1.5mm i.d.) were used, and the stationary liquid phases (SF-96–QF-1, 3:1) were supported on GasChrom G HP AW DMCS (100–120 mesh). The capillary column (30m × 4mm i.d.) was coated with OV-101 by a dynamic procedure [149]. Unless the temperature of the column oven was programmed, the temperatures of the injector, column and detector were 225, 185 and 220°C, respectively. With packed columns the flow rate of the carrier gas (nitrogen) was adjusted to ca. 20mL min-1. With capillary columns a minimum flow rate of 1.9mL min-1 was required to operate the electron-capture detector. The flows of hydrogen and air to the flame ionization detector were ca. 25 and 250mL min -1, respectively. To prepare the detection system for open tube capillary columns, a low volume detector was developed. The electron-capture detector is sensitive to solute concentration, and the use of a scavenging gas dilutes the sample and lowers the sensitivity. By modification of the arrangement of the tritium foil in the detector cell (Fig. 9.6), a cell volume of 0.15mL was obtained. The detector was operated with comparable sensitivity to that of a conventional cell at a carrier gas flow rate of 1.9mL min-1.
528
Insecticides and herbicides
Fig. 9.6 Modified arrangement of the tritium foil in the electron-capture detector cell. (Reprinted with permission from Södergren [147]. Copyright (1978) Elsevier Science Publishers BV.)
The gas chromatograms in Fig. 9.7 show that organochlorine insecticides and methyl esters of fatty acids were detected simultaneously using this system. Bacaloni et al. [150] used capillary column gas chromatography for the analysis of mixtures of chlorinated insecticides, PCBs and other pollutants. Graphitized capillary columns were used in this work. These columns have the interesting feature that according to the amount of stationary phase coated on the walls, they operate in gas–solid, gas– liquid and gas–liquid– solid chromatographic modes. Bacaloni et al. [150] used columns coated with PEG 20M and it is shown that selective columns can be obtained that give different performances for particular applications. Bacaloni et al. [150] gas chromatographed mixtures of 15 chlorinated pesticides on columns of different lengths and loadings of stationary phase under the same operation conditions at 160°C. Bacaloni et al. [150] concluded that a glass capillary column loaded with a large amount of PEG 20M is suitable for the analysis of volatile chlorinated compounds, phenols and amines, whereas for the analysis of chlorinated compounds, including the PCBs, a column with a low loading of stationary phase is desirable. On the basis of these results, the separation of mixtures of chlorinated insecticides and PCBs (Arochlor 1242 and 1260) was tried on a 70m column having 210 000 theoretical plates for hexadecane at 100°C and 160 000 plates for ?-BHC at 160°C. Figure 9.8 shows chromatograms of complex mixtures of chlorinated organic compounds. The Department of the Environment (UK) [130] has used a tentative method for the determination of organochlorine insecticides and PCBs
Mixtures of chlorinated insecticides and polychlorobiphenyls
529
Fig. 9.7 Simultaneous response of electron-capture detector–flame ionization detector (ECD– FID) to a mixture of organochlorine insecticides and methyl esters and fatty acids. Peaks: 1 = Lindane; 2 = BHC; 3 = Aldrin; 4 = p,p’-DDE; 5 = Dieldrin; 6 = p,p’-DDD; 7 = p,p’-DDT; 9 = myristic acid; 10 = palmitic acid; 11 = stearic acid; 12 = oleic acid. (Reprinted with permission from Södergren [147]. Copyright (1978) Elsevier Science Publishers BV.)
in natural and drinking waters and sewage effluents. The first part of this method is concerned with the extraction and determination of the amounts of individual substances present; the second part is concerned with methods for verifying the identity of the various substances quantified in the first part. Representative data are presented for a number of typical insecticides using different chromatographic stationary phases (Table 9.12). Information on blanks and limits of detection are given in Table 9.13. Millar et al. [17] have described a gas chromatographic method, utilizing an electron-capture detector, for the determination of 25 organochlorine insecticides and PCBs in water. Data are presented on extractability, recovery from clean-up columns, recovery from spiked waste waters, and preservation of samples for analysis. Thomas [151] has described a dual-column, dual-detection gas chromatograph system for the determination of chlorinated insecticides and PCBs in environmental waters. Leoni [152] separated 50 organochlorine insecticides and PCBs into four groups by silica gel multicolumn chromatography. The separation was undertaken to simplify the chromatograms obtained by gas
Fig. 9.8 Gas chromatograms of mixtures of pesticides and Arochlors on column 5. Carrier gas, H ; electron2 capture detector. Numbers of peaks correspond as follows: 1 α-BHC, 2 Heptachlor, 3 Aldrin, 4 α-BHC, 5 α-Endosulphan, 6 α-Chlordane, 7 α-Chlordane, 8 o,p’-DDE, 9 Dieldrin, 10 Endrin, 11 p,p’-DDE, 12 Perthane, 13 o,p’-DDD, 14 Endosulphan, 15 p,p’-DDT. (a) Arochlor 1242; temperature programme, 140°C for 95min, then increased from 140 to 160° at 2°C min-1; *interfering peaks. (b) Arochlor 1254; temperature programme 140°C for 105min then increased from 140 to 160°C at 2°C min-1. Alternative programme: 140°C for 170min then increased from 140 to 160°C at 2°C min-1 (see inset peaks); *interfering peaks, (c) Arochlor 1260: temperature programme, 140°C for 185min, then increased from 140 to 160°C at 2°C min-1. Alternative programme; isothermal at 160°C (see inset peaks): b–f are peaks of Arochlor.
Mixtures of chlorinated insecticides and polychlorobiphenyls
531
Table 9.12 Retention lines of organochlorine insecticides relative to Dieldrin, on some GLC columns
Note a Chlordane is a multipeak compound with other smaller peaks at RRT 0.37, 0.69, 0.76 and 1.23. Glass columns 1m × 3mm internal diameter 60–80 mesh acid-washed DMCS treated Chromosorb W support at l80°Cand20–40ml nitrogen min-1.
chromatography from contaminated samples. The sample (10–15L of surface water or 20–25L of potable water) was acidified with hydrochloric acid to pH3, and subjected to continuous extraction, twice with light petroleum (boiling range 40–60°C) and once with benzene. The benzene extract was evaporated and the residue was dissolved in light petroleum and added to the other extracts. The combined extracts were concentrated to 15mL and were partially purified by extraction with acetonitrile saturated with light petroleum (4 × 30mL). The extract was diluted with 2% sodium chloride solution (700mL) and the solution was again extracted with light petroleum (2 × 100mL). This extract was evaporated and the residue was dissolved in 1mL of hexane. This solution, followed by 1mL of hexane used for rinsing the container, was
532
Insecticides and herbicides
Table 9.13 Blanks and limits of detection
Notes Sw = within batch standard deviation. Sb = between batch standard deviation. St = total standard deviation.
applied to a column (100mm × 4.2mm) of silica gel (Grace 950, 60– 200 mesh, dried at 30°C for 2h then deactivated with 5% of water). The column was then percolated (at 1mL min-1) in turn with hexane (20mL), benzene–hexane (3:2) (8mL), benzene (8mL) and ethyl acetate–benzene (1:1) (14mL). Each eluate was evaporated and the residue dissolved in 1mL of hexane and subjected to gas chromatography on OV-17 as stationary phase with electron-capture detection. Quantitative recovery was achieved for all insecticides except Malathion, Disulfoton, Dimethoate and Phorate. 9.2.1.2 Supercritical fluid chromatography Bavel et al. [153] have developed a solid-phase carbon trap for the simultaneous determination of polychloro insecticides, PCBs, PCDDs and PCDFs in natural waters using supercritical fluid extraction liquid chromatography. Recoveries were in the range 57– 134%. 9.2.1.3 Voltammetry Farwell et al. [154] used voltammetry to identify organochlorine insecticides, PCBs, polychlorinated naphthalenes, and polychlorinated benzenes. They list tables of reduction potentials for over 100 organochlorine compounds.
Mixtures of chlorinated insecticides and polychlorobiphenyls
533
9.2.1.4 Miscellaneous Millar et al. [17] carried out experiments to study a method for the recovery of 18 organochlorine pesticides and seven PCBs from water. Extractions with dichloromethane, and 15% dichloromethane in hexane, at pH2, 7 and 10 and liquid–solid column chromatography using columns of Florasil or alumina produced excellent results. An investigation was also made into the effects of pH, temperature, and residual chlorine on the preservation of spiked samples and recommendations are made for the most suitable storage conditions. 9.2.1.5 Preconcentration Leoni et al. [155, 156] observed that in the extraction of organochlorine insecticides and PCBs from surface and coastal waters in the presence of other pollutants such as oil, surface active substances etc., the results obtained with an absorption column of Tenax–Celite (a porous polymer; trademark registered by Enka NV; developed by AKZO Research Labs, Arnhem, Netherlands) are equivalent to those obtained with continuous liquid–liquid extraction technique. For natural waters that contain solids in suspension that absorb pesticides, it may be necessary to filter the water before extraction with Tenax and then to extract the suspended solids separately. Analyses of river and estuarine sea waters, filtered before extraction, showed the effectiveness of Tenax, and the extracts obtained for organochlorine and organophosphorus pesticide analysis prove to be much less contaminated by the liquid–liquid technique. Leoni et al. [156] showed that for the extraction of organic micropollutants such as pesticides and aromatic polycyclic hydrocarbons from waters, the recoveries of these substances from unpolluted waters when added at the level of 1ppb averaged 90%. Leoni et al. [155, 156] conclude that the extraction of insecticides from waters by adsorption on Tenax yields results equivalent to those by the liquid-liquid procedure when applied to mineral, drinking and surface waters that completely or almost completely lack solid matter in suspension. For waters that contain suspended solids that can adsorb some insecticides in considerable amounts, the results of the two methods are equivalent only if the water has previously been filtered. In these instances, therefore, the analysis will involve filtered water as well as the residue of filtration. Compared with liquid–liquid extraction, the main advantages of Tenax are the considerable amount of time saved, the possible automation of the process and the fact that gas chromatographic analysis shows the ‘extracts’ obtained with Tenax to be less contaminated by interfering substances. Another advantage of Tenax is that the product can be used ‘as received’, without preliminary treatment.
534
Insecticides and herbicides
Gosink [157] and Aue [158] have also studied the determination of organochlorine insecticides and PCBs in water samples. Sackmauereva et al. [14] used columns filled with silicic acid–Celite to separate organochlorine insecticides from PCBs. The PCBs were eluted with petroleum ether. To elute insecticides from the column they used a mixture of acetonitrile and hexane and methylene chloride. C -silica-bonded phase columns [92, 159] have been used to 18 preconcentrate chlorinated insecticides prior to gas chromatographic [159] and thin-layer chromatographic analysis [92]. Noy et al. [160] have discussed an on-line liquid chromatography–gas chromatography combination for the preconcentration of chlorinated insecticides, Noroozian et al. [94] have described a sample on-line technique involving sorption on a liquid chromatography micro precolumn packed with reversed-phase adsorbent followed by direct elution gas chromatography with hexane to determine a series of chlorinated insecticides and polychlorinated biphenyls in natural waters. Amberlite XAD-2 resin [89, 161–163], cellulose triacetate membrane filters [164] and Tenax polymer [155] have all been used to obtain concentrated extracts of chlorinated insecticides and PCBs from water samples. 9.2.2 Sea water 9.2.2.1 Gas chromatography Girenko et al. [165] noted that it was difficult to analyse samples of sea water because they are severely polluted by various coextractive substances, chiefly chlorinated biphenyls. To determine organochlorine insecticide residues by gas chromatography with an electron-capture detector, the chlorinated biphenyls were eluted from the column together with the insecticides. They produce inseparable peaks with equal retention times, thus interfering with the identification and quantitative determination of the organochlorine insecticides. The presence of chlorinated biphenyls is indicated by additional peaks on the chromatographs of the water samples and aquatic organic organisms. Some of the peaks coincide with the peaks of the o,p’ and p,p’ isomers of DDE, DDD and DDT and some of the constituents are eluted after p,p’-DDT. 9.2.2.2 Preconcentration Organochlorine insecticides and PCBs have been preconcentrated from sea water on to a column of Tenax–Celite as discussed in Section 9.2.1.5 [155, 156].
Mixtures of chlorinated insecticides and polychlorobiphenyls
535
9.2.3 Potable waters 9.2.3.1 Gas chromatography The gas chromatographic method described by the Department of the Environment [130] discussed in Section 9.1.4.1 has also been applied to the analysis of chlorinated insecticides and PCBs in potable water. 9.2.4 Waste waters 9.2.4.1 Gas chromatography A method has been described [166] for the determination of the following PCBs (Arochlors) at the nanogram level in 15:85 methylene chloride– hexane extracts of water and waste water: PCB-1016, PCB-1221, PCB1232, PCB-1242, PCB-1248, PCB-1254 and PCB-1260. This method is an extension of the method for chlorinated hydrocarbons in water and waste water (described by Goerlitz and Law [167]). It is designed so that determination of both PCBs and the following organochlorine insecticides can be made on the same sample: Aldrin, DDT, Mirex, BHC, Heptachlor, pentachlorinitrobenzene, Chlordane, Heptachlor epoxide, Strobane, DDD, Lindane, Toxaphene, DDE, Methoxychlor and Trifluralin. The PCBs and the organochlorine insecticides are coextracted by liquid– liquid extraction and insofar as possible, the two classes of compounds are separated from one another prior to gas chromatographic determination. A combination of the standard Florasil column clean-up procedure and a silica gel microcolumn separation procedure are employed. Identification is made from gas chromatographic patterns obtained through the use of two or more unlike columns including SE-30 or OV-1 (3%), OV-17 (1.5%) or OV-210 (1.95%) on Gas Chrom. Detection is accomplished using an electron-capture, microcoulometric or electrolytic conductivity detector. Techniques for confirming qualitative identification are suggested by these workers. The interference in industrial effluents is high and varied and poses great difficulty in obtaining accurate and precise measurement of PCBs and organochlorine insecticides. Separation and clean-up procedures are generally required and may result in the loss of certain organochlorine compounds. Therefore, great care should be exercised in the selection and use of methods for eliminating or minimizing interferences. Phthalate esters, certain organophosphorus insecticides and elemental sulphur will interfere when the microcoulometric or electrolytic conductivity detectors are used in the halogen mode. Organochlorine insecticides and other halogenated compounds constitute interferences in the determination of PCBs. Most of these are separated by the method described. However,
536
Insecticides and herbicides
certain compounds, if present in the sample, will occur with the PCBs. Included are: sulphur, Heptachlor, Aldrin, DDE, Chlordane and Mirex. Workers at the US Environmental Protection Agency [168] have evaluated protocols for chlorinated insecticides and PCBs in raw waste water. They concluded that the gas chromatographic method performed satisfactorily at the parts per billion level. Millar et al. [17] have described a gas chromatographic method for the determination of 25 organochlorine insecticides and PCBs in waste water. Data are presented on extractability, recovery from clean-up columns, and preservation of samples for analysis. McIntyre et al. [169] determined organochlorine pesticides and PCBs by electron-capture gas chromatography. The waste water sample is extracted with hexane and the extract cleaned up on alumina–silver nitrate and silica gel columns prior to electron-capture gas chromatography. Dunn et al. [170] used gas chromatography data to carry out pattern recognition for classification and determination of PCBs in waste waters and waste dump extracts. Ericksen et al. [171] used high-resolution gas chromatography– electron-impact mass spectrometry to determine PCBs in halocarbon solvent process and product wastes. 9.2.5 Sewage effluents 9.2.5.1 Gas chromatography Mattson and Nygren [172] devised a solvent extraction method for extracting PCBs from sewage sludge containing lipids. They point out that lipids and some other impurities in the crude extracts of sewage sludge can be destroyed by treatment with fuming sulphuric acid, either by shaking the acid [173] or by eluting on a fuming sulphuric acid-Celite column [174, 175]. Dieldrin is decomposed by this treatment but DDT and its metabolites, DDD and DDE, are not. Extracts of sewage sludges often contain large amounts of elemental sulphur, particularly after treatment with sulphuric acid. These interfere with early eluting compounds in the gas chromatographic step. Sulphur was removed by the Bartlett and Skoog [176] method in which the sulphur is reacted with cyanide in acetone solution to produce thiocyanate. BHCs are decomposed to some extent, probably to pentachlorocyclohexane. An alternative procedure for the removal of sulphur utilizing barium hydroxide is also described. Alkali hydroxides should not be used as they cause dehydrochlorination of BHCs [173]. Lindane and its isomers are dehydrochlorinated to trichlorobenzenes [177] and are eluted together with the solvent. Cochrane and Maybury [178] have used the reaction with sodium hydroxide in methanol for the
Mixtures of chlorinated insecticides and polychlorobiphenyls
537
identification of BHCs. Dieldrin is not decomposed in the potassium hydroxide treatment and can thus be detected in the chromatogram of that aliquot. Some common chlorinated hydrocarbon pollutants and the internal standard hexabromobenzene were treated, according to the general procedure described above with sulphuric acid, potassium cyanide and potassium hydroxide. The results of the recovery experiments are shown in Table 9.14. When using packed columns, a precolumn of sodium and potassium hydroxides will give the same effect as the potassium hydroxide treatment described above [179]. Mattson and Nygren [172] have also tested a column with a packed alkaline postcolumn to remove the sulphur peak from the chromatogram. In the postcolumn, DDT and DDD are dehydrochlorinated but this does not affect their retention times. This method has good reproducibility and has a detection limit for the total amount of PCBs in the dried sample of at least 0.1mg kg-1 and for DDT, DDD and DDE limits of 0.01, 0.005, and 0.005mg kg-1, respectively. Jensen et al. [180] have described a procedure for the determination of organochlorine compounds including PCBs and DDT in sediments and sewage sludge in the presence of elemental sulphur. The method can also be used for a search for both volatile and/or polar pollutants. The sulphur interfering in the gas chromatographic determination is removed in a nondestructive treatment of the extract with tetraburylammonium sulphite. Table 9.14 Effect of treatment of a solution of chlorinated hydrocarbons and the internal standard hexabromobenzene with fuming sulphuric acid (I), fuming sulphuric acid plus potassium cyanide (II), and potassium hydroxide (III) expressed as percentages of the compounds in an untreated solution
Source: Reprinted with permission from Cochrane and Maybury [178]. Copyright (1973) American Chemical Society.
538
Insecticides and herbicides
This lipophilic ion pair rapidly converts the sulphur to thiosulphate in an organic phase. The recovery of added organochlorines was above 80% and the detection limit in the range of 1–10ppb organochlorines from a 10g sample. Elemental sulphur present in most sediment and digested sludge has caused significant problems in residue analysis [55, 181]. If the sulphur level is high, the electron-capture detector will be saturated for a considerable period of time, and if the level of sulphur is low, it gives three or more distinct peaks on the chromatogram which can interfere with BHC isomers and Aldrin. Treatment of the crude extract with potassium hydroxide in ethanol [173] or Raney nickel [182] will quantitatively destroy all sulphur, but will at the same time convert DOT and DDD to DDE and DDMU (1-chloro-2,2-bis(4-chlorophenyl)ethane), respectively, and most BHC isomers are lost. Metallic mercury has also been used for removal of sulphur [183]. Jensen et al. [180] described an efficient, rapid, nondestructive method to remove the sulphur according to the reaction
where TBA+ is the tetrabutylammonium ion. The sample extracts were run with an electron-capture detector (63Ni). The 2.4m × 0.18cm (i.d.) glass column was filled with a mechanical mixture of two parts of 8% QF-1 and one part of 4% SF-96 on acidwashed silanized Chromosorb W (100–120 mesh). Typical chromatograms obtained by this procedure are shown in Fig. 9.9. The gas chromatographic method described by the Department of the Environment [130] discussed in Section 9.1.4.1 has also been applied to the analysis of chlorinated insecticides and PCBs in sewage effluents. Workers at the US Environmental Protection Agency [168] have evaluated protocols for chlorinated insecticides and PCBs in raw waste water and sewage effluents. They concluded that the gas chromatographic method performed satisfactorily at the parts per million level. McIntyre et al. [184,185] described a method for the analysis of PCBs and chlorinated insecticides in sewage sludges in which homogenized samples are extracted with hexane, concentrated and cleaned up on an aluminaalumina plus silver nitrate column and eluted with hexane. After concentration of the eluent, PCBs and organochlorine compounds were determined by a silica gel chromatographic procedure and gas chromatography. The gas chromatograph was fitted with a 63Ni electron-capture detector. Glass columns (2cm × 3mm i.d.) containing 1.5% OV-17 plus 1.95% QF-1 on 100–120 mesh Supelcoport were used for primary identification and 1% NPGG on 80–100 mesh Gas Chrom Q for confirmation. McIntyre et al. [184, 185] found that the method by Holden and Marsden [186] resulted in good reproducible separation of PCBs and
Mixtures of chlorinated insecticides and polychlorobiphenyls
539
Fig. 9.9 (a) Typical digested sewage sludge chromatogram, severely contaminated with sulphur, (b) The same sample after a normal TBA-sulphite treatment, showing that most of the sulphur has disappeared. A number of peaks (a, b, c) originating from traces of sulphur appear in the BHC–Aldrin region, (c) The final chromatogram after additional treatment with sodium sulphite. • = PCB components, is = internal standard. (Reprinted with permission from Jensen et al. [180]. Copyright (1977) American Chemical Society.)
540
Insecticides and herbicides
organochlorine insecticides. The results indicate that recovery from four compounds was quantitative with recoveries of 99.8, 99.6, 102.5 and 102.3% for Arochlor 1260, p,p’-DDE, γ-HCH, and Dieldrin, and displays little scatter, with relative standard deviations of 1.6, 3.1, 2.9 and 3.1% respectively for the four determinations. 9.2.5.2 Gas chromatography–mass spectrometry Erikson and Pellizzari [187] analysed municipal sewage samples in the USA by a gas chromatography–mass spectrometry–computer technique for chlorinated insecticides and PCBs. The samples (300g) were extracted at pH11 six times with a total of 350mL chloroform to remove neutral and basic compounds. The extract was dried with sodium sulphate, vacuum filtered, and concentrated to 2mL using Kuderna–Danish apparatus. In cases where the sample background interfered significantly, an aliquot of the sample was chromatographed on a 1.0 × 30cm silica gel column (Snyder and Reinert [188]). PCBs and related compounds were eluted with 50mL hexane; pesticides and other compounds were eluted with 50mL toluene. Acidic components of the samples were treated as diazomethane and dimethyl sulphate (Keith [189–191]). Analysis of all samples for PCBs was accomplished using a Finnigan 3300 quadruple gas chromatograph–mass spectometer with a PDP/12 computer. The 180cm × 2mm i.d. glass column, packed with 2% V-101 on Chromosorb W, was held at 120°C for 3min, programmed to 230°C at 12°C min-1 and held isothermally until all peaks had eluted. Helium flow was 30mL min-1. The ionization voltage was nominally 70eV and multiplier voltages were between 1.8 and 2.2kV. Full scan spectra were obtained from m/e 100–500. Samples were analysed under the following temperature conditions: up to 150°C for 3min, programmed to 230°C at 80min1 isothermally until all peaks had eluted. PCBs were quantitated by a gas chromatography–mass spectrometry– computer technique using the selected ion-monitoring mode to provide maximum sensitivity and precision. This technique has been used in similar work on polychlorinated naphthalenes (Erikson et al. [192, 193]). Ten ions were selected for monitoring: one from the parent cluster for each of the chlorinated biphenyls (C12H9Cl to C12Cl10). PCBs were quantitated using anthracene as external standard and a previously determined relative molar response (anthracene parent ion mass 178; 27ng mL1 ). Anthracene does not interfere with PCB determination, nor do PCBs or their fragment ions interfere with the determination of anthracene. The retention time results for 34 chlorinated compounds found in sewage are given in Table 9.15. Not all compounds could be identified. A large number of spectra contained what appeared to be chlorine isotope clusters, which are not reported. This could be due to interferences, very low levels, or spurious peaks. Although no structure could e assigned, the mass spectra indicated possible structures for three
Mixtures of chlorinated insecticides and polychlorobiphenyls
Table 9.15 Summary of chlorinated compounds found in sewage sludge
Source: Reprinted with permission from Erikson and Pellizzari [187]. Copyright (1979) Springer-Verlag, New York. Notes a Unidentified compounds are listed with the apparent molecular weight and number of chlorines. If the identification of a compound is tentative, it is denoted by (tent.). b Retention times are listed for the chromatographic temperature conditions, 12°C for 3min, then 12° min-1 to 230°C, then hold. Values in parentheses are for chromatographic temperature conditions, 150°C for 3min, then 8° min-1 to 230°C, then hold. c Two separate isomers observed in some samples. d Differences in retention times possibly indicate different isomers.
541
542
Insecticides and herbicides
compounds in Table 9.15. The compound containing two chlorines with mol wt = 187 (RT = 2.3–3.7min) may have the molecular formula C H Cl 8 7 2 and could be a dichlorodihydroindole or related compounds. Two distinct compounds were observed with four chlorines mol wt = 240. These compounds appear to be isomers of tetrachlorostyrene (C H Cl ). 8 4 4 The 4,4'-dichlorobenzophenone identification was confirmed by comparison of the retention time with an authentic sample. The two peaks identified as DDE isomers are probably the two common isomers, o,p’-DDE and p,p’-DDE, which generally are separable by gas chromatography. Examination of neutral extracts of sewage sludge by gas chromatography–mass spectrometry revealed the presence of appreciable amounts of PCBs. 9.2.5.3 Miscellaneous Various other workers [194–200] have reported methods for the determination of PCBs and organochlorine pesticides in sewage and sewage sludges. 9.3 Phosphorus-containing insecticides 9.3.1 Natural waters 9.3.1.1 Spectrophotometric methods Venkataraman and Sathyamurthy [201] developed a simple and direct spectrophotometric method for the determination of Parathion in water in which a benzene extract of Parathion is hydrolysed and is reduced with zinc dust in acid solution. The resulting amino derivative is diazotized with sodium nitrite and hydrochloric acid. The diazo compound is coupled with a naphthylethylenediamine hydrochloride and a magenta dye is produced, which is evaluated spectrophotometrically. It is unlikely that this method is specific for Parathion. Glass has described a spectrophotometric method [202] for the determination of glyphosphate in natural water. Frobe et al. [203] compared hydrolytic degradation and oxygen combustion methods of decomposition for the organophosphorus pesticides, Malathion, Parathion and Phosalone. The pesticides were decomposed to orthophosphoric acid prior to spectrophotometric determination as phosphomolybdenium blue. Pesticide adsorption on Amberlite XAD-4 resin prior to combustion increased the sensitivity of the method while eliminating both inorganic phosphates and some nonpesticide organophosphorus compounds. A tandem approach involving
Phosphorus-containing insecticides
543
the determination of total organophosphorus pesticides and a test for chlorinesterase inhibitors was recommended for differentiating between various sources of phosphorus; an increased phosphorus content and a positive cholinesterase inhibition test would clearly indicate the presence of organophosphorus pesticides in surface waters. 9.3.1.2 Flow injection analysis Burguera and Burguera [204] give details of equipment and a procedure for the determination of several organophosphorus insecticides (Dicrotophos, Dimethoate, Malathion and Parathion) in admixture in water by flow injection analysis combined with molecular emission cavity analysis. The insecticides were extracted from the sample with hexane and dichloromethane at pH7.0, and injected into the flow system. Recoveries were 73.4–98.1% for insecticide concentrations containing 50g phosphorus. 9.3.1.3 Gas chromatography Much of the development work on the determination of organophosphorus insecticides which has been carried out in recent years has hinged on the development of suitable detectors which are ultra-sensitive and which are specific for phosphorus in the presence of other elements such as carbon, hydrogen, oxygen, halogens, nitrogen and sulphur and indeed in some cases, can be used to determine compounds containing these other elements. Several types of organophosphorus insecticides also contain halogens, nitrogen, or sulphur. Electron-capture, flame ionization, flame photometric, microcoulometric, thermionic and electrolytic conductivity detectors have all been studied in this application (Table 9.16). The advantages and disadvantages of these types of detectors are discussed below in general proceeding to a discussion of the application of gas chromatography to the determination of particular organophosphorus insecticides, or mixtures thereof, in water. The quantitative analysis of organophosphorus pesticide residues has become much easier since the development of the alkali flame and flame photometric detectors, both of which are now commercially available. The Karmen–Guiffrida detector (the so-called thermionic detector) has proved extremely useful. It is basically a flame ionization detector with an alkali salt ring placed on the flame tip. Not only does this enhance the sensitivity but also the selectivity. This may be of the order of 104–105 for phosphorus-containing compounds as compared with the equivalent carbon compounds. The selectivity for halides and nitrogen is between 102and 103 and it is 102 for sulphur and about 10 for arsenic. The response for phosphorus increases with increasing hydrogen flow but some increase in the background current is also observed. A variety
544
Insecticides and herbicides
Table 9.16 Detection used in gas chromatography of phosphorus-containing organic compounds
Phosphorus-containing insecticides
545
of alkali salts are used including potassium chloride, caesium bromide and rubidium sulphate. The type and shape of the tip and anode, together with the flow rates, are very much a matter of personal choice. De Loach and Hemphill [205, 206] have discussed the design of a rubidium sulphate detector and with optimum conditions claim a sensitivity of 1pg. Often a charcoal column clean-up is used before injection to prevent the column becoming contaminated. Brazhnikov et al. [207] developed a highly stable thermionic detector (the thermaerosol detector) to avoid the limitations imposed by modifications of existing detectors. The thermaerosol detector, which combines a conventional flame ionization detector with a generator of an aerosol of alkali metal salt was applied to the analysis of organophosphorus insecticides. The detector avoids limitations of conventional thermionic detectors such as a considerable dependence of
546
Insecticides and herbicides
the sensitivity and reproducibility of the detector operation on the flow rate of hydrogen, air and carrier gas, rapid exhaustion of the alkali metal salt source and the difficulty of replacing one salt by another. The important advantages of this detector are the simplicity of its design, the possibility of rapid replacement of the salt without dismantling the detector and stability of its operation for a long period (several thousand hours) without the need to replace the salt reservoirs. The most commonly used organophosphorus insecticides are listed below in alphabetical order. From this list can be seen the wide variety of these phosphorus-based compounds in use. Gas chromatography has played a major part in the development of suitable methods for the determination of submicro amounts of such substances in environmental samples, such as crops, animal tissues and water. Abate Acciothion (Sumithion, Fenitrothion, Folithion) Amidithion
Amiton Azinphos-ethyl
Azinphos-methyl
Azothoate
Bensulide
Bromophos
Bromophos-ethyl
Butonate
O,O,O’,O’-tetramethyl-O,O’-(thiodi-p-phenylene diphosphorothioate) O,O-dimethyl-O-4-nitro-m-ethyl phosphorothioate S-(N-2-methoxyethylcarbamoylmethyl)dimethyl phosphorothiolothionate S-(N-2methoxyethylcarbamoylmethyl)-O,O-dimethyl phosphorodithioate S-(2-(diethylamino)ethyl)-O O-diethyl , phosphorothionate S-(3,4-dihydro-4-oxobenzo(d)-[1,2,3]triazin-3ylmethyl) diethyl phosphorothiolothionate S-(3,4dihydro-4-oxobenzo(d)-[1,2,3]triazin-3-ylmethyl)O,O-diethyl phosphorodithioate S-(3,3-dihydro-4-oxobenzo(d)-[1,2,3]triazin-3ylmethyl)-dimethyl phosphorothiolothionate S-(3,4dihydro-4-oxobenzo(d)-[1,2,3]triazin-3-ylmethyl)O,O-dimethyl phosphorodithioate 4-(4-chlorophenylazo)phenyldimethyl phosphorothionate O-4-(4-chlorophenylazo)phenylO O-dimethyl phosphorothioate , N-2-(O,O-di-isopropylphosphorothiolothioyl)ethyl benzenesulphonamide di-isopropyl S-(2phenylsulphonylaminoethyl)phosphorothiothionate O,O-di-isopropyl S-(2phenylsulphonylamino-ethyl)-phosphorodithioate 4-bromo-2,5-dichlorophenyldimethyl phosphorothionate O-(4-bromo-2,5-dichlorophenyl)O,O-dimethyl phosphorothioate (4-bromo-2,5-dichlorophenyldiethyl) phosphorothionate O-(4-bromo-2,5-dichlorophenyl)O,O-diethyl phosphorothioate dimethyl-1-butyryloxy-2,2,2-
Phosphorus-containing insecticides
Carbophenothion
Chlorfenvinphos Chlorphonium Chlorphoxim
Coumaphos
Coumithoate
Crotoxyphos
Cyanophos Dasanite Demephion Demephion-O
Demephion-S
Demeton Demeton-methyl Demeton-O Demeton-O-methyl Demeton-S Demeton-S-methyl
547
dichloroethylphosphonate S-(4-chlorophenylthiomethyl)diethyl phosphorothiolothionate S-(4chlorophenylthiomethyl)-O,O-diemethyl phosphorodithioate 2-chloro-1-(2,4-dichlorophenyl)vinyldiethyl phosphorodithioate tributyl-2,4-dichlorobenzylphosphonium (2-chloro-a-cyanobenzylideneamino)diethyl phosphorothionate O-2chlorocyanobenzylideneamino-O,O-diethyl phosphorothioate 2-chloro-a(diethoxyphosphinothioyloxyimino)phenylacetonitrile 3-chloro-4-methyl-7-coumarinyldiethyl phosphorothionate O-(3-chloro-4-methyl-7coumarinyl)-O,O-diethyl phosphorothionate diethyl-7,8,910,-tetrahydro-6-oxobenzo(c)criroman3-yl phosphorothionate O,O-diethly-O-(7,8,9,10tetrahydro-6-oxobenzo(c)chroman-3-yl) phosphorothioate O,O-diethly-O-(7,8,9,10tetrahydro-6-oxo-6didibenzo(bd)pyran-3-yl) phosphorothionate dimethyl cis-1-methyl-2-(1-phenylethoxycarbonyl) vinyl phosphate 1-methylbenzyl-3(dimethyoxyphosphinyloxy) isocrotonate 4-cyanophenyl phosphorothionate O-4cyanophenyl-O,O-dimethyl phosphorothionate O,O-diethyl-O-(4-methylsulphinylphenyl) phosphorothioate mixture of Demephion-O and Demephion-S (see below) dimethyl-2-(methylthio)ethyl phosphorothionate O,O-dimethyl-O-(2-(methylthio)ethyl) phosphorothioate dimethyl-S-(2-(methylthio)ethyl) phosphorothiolate O,O-dimethyl-S-(2(methylthio)ethyl phosphorothioate A mixture of Demeton-O and Demeton-S (see below) A mixture of Demeton-O-methyl and Demeton-Smethyl (see below) diethyl-2-(ethylthio)ethyl phosphorothionate O,Odiethyl-O-(2-(ethylthio)ethyl) phosphorothioate 2-(ethylthio)ethyldimethyl phosphorothionate O-(2(ethylthio)ethyl-O,O-dimethyl phosphorothioate diethyl-S-(2-(ethylthio)ethyl) phosphorothiolate O,O-diethyl-S-(2-(ethylthio)ethyl) phosphorothioate S-(2-(ethylthio)ethyl)dimethyl phosphorothiolate S-
548
Insecticides and herbicides
Diazinon
Dichlorvos Disulfoton Dursban (Chlorpyrifos) Edifenphos Ethion Fenchlorphos (Ronnel) Fenthion Fonofos GS-13005 Methocrotophos
Monitor Nemacur Paraoxon Parathion Parathion-ethyl Phorate Pirimiphos-ethyl
Pirimiphos-methyl
Prothoate Salithion Tetrachlorvinphos Velsicol VCS-506
(2-(ethylthio)ethyl)-O,O-dimethyl phosphorothioate diethyl-2-isopropyl-6-methyl-4-pyrimidinyl phosphorothionate O,O-diethyl-O-(2-isopropyl-6methyl-4-pyrimidyl) phosphorothioate 2,2-dichlorvinyldimethyl phosphate O,O-dimethyl-S-(2-(ethylthio)ethyl) phosphorodithioate O,O-dimethyl-O-(3,5,6-(triethylchloro-2-pyridyl) phosphorothioate ethyl-S,S-diphenyl phosphorodithiolate thionate Oethyl-S,S-diphenyl phosphordithioate S,S-methylene-O,O,O’,O’-tetraethyl phosphorodithioate O,O-dimethyl-O-2,4,5-trichlorphenyl phosphorothioate O,O-dimethyl-O-(4(methylthio)-m-tolyl) phosphorothioate ethyl-S-phenylethyl phosphorothiolothionate ethylS-phenylethyl phosphorodithioate S-(5-methoxy-2-oxo-1,3,4-thiadazolin-3-yl)-O,Odimethyl phosphorodithioate dimethyl-cis-2-(N-methoxy-N-methyl-carbamoyl)-1methylvinyl phosphate 3(dimethyloxyphosphinyloxy)-N-methoxy-Nmethylisocrotonamide O,S-dimethyl phosphoramidothioate ethyl-4-(methylthio)-m-tolyl isopropyl phosphoramidate The oxygen analogue of Parathion O-(4-nitrophenyl phosphorothioate) O,O-dimethyl-S-(ethylthio)methyl phosphorodithioate 2-diethylamino-6-methypyrimidin-4-yl diethyl phosphorothionate O-(2-diethylamino-6methylpyrimidin-4-yl)-O,O-diethyl phosphorothioate 2-diethylamino-6-methylpyrimidin-4-yl-dimethyl phosphorothionate O-(2-diethylamino-6methylpyrimidin-1-yl)-O,O-dimethyl phosphorothioate O,O-dimethyl-S-N-isopropyl carbamoyl methyl phosphorothiolothionate 2-sulphide of 2-methoxy-4-H-benzo-1,2,3dioxaphosphrin trans-2-chloro-1-(2,4,5trichlorophenyl)vinyldimethyl phosphate O-(4-bromo-2,5-diethylchorophenyl)-Omethylphenyl phosphorothioate
Phosphorus-containing insecticides
Zyron
549
O-2,4-dichlorophenyl-O-methylisopropyl phosphoramidothioate
Askew et al. [53] have developed an early general method for the determination of organophosphorus insecticide residues and their metabolites in river waters and sewage effluents utilizing gas chromatography. The organophosphorus pesticides vary greatly in their polarity, and the extent of their extraction from aqueous samples is markedly dependent on the nature of the solvent used. Table 9.17 gives a comparison of the efficacies of three solvents in removing these insecticides from water, and it is apparent that a polar solvent such as chloroform is the most generally used. Askew et al. [53] found that most water extracts are sufficiently low in coextractive to require no clean-up for gas chromatographic purposes. The nature of the coextractive will undoubtedly vary with the location from which the samples are taken, and no complete comprehensive clean-up procedure can be recommended. A column containing 1g of Nuchar carbon eluted with chloroform gave the most useful clean-up where this was found to be necessary. The following insecticides are exceptional in being retained by the column described: Azinophos-ethyl, Azinophos-methyl, Coumaphos, Dichlorvos, Maloxon, Menazon, Phosalone and Vamidothion. When these insecticides are encountered a clean-up on alumina [208] or magnesium oxide [209] would be preferable. The general screening technique developed by Askew et al. [53] is described in full in Fig. 9.10. Stage (a) is applied as a general screening technique, and stage (b) is incorporated only when suspected pesticides are encountered. Figure 9.11 shows gas chromatographs obtained by Askew et al. [53] on solvent extracts of River Thames water unspiked and spiked Table 9.17 Extraction of insecticides from aqueous solution by organic solvents
Source: Reprinted with permission from Askew et al. [53]. Copyright (1969) Royal Society of Chemistry. Note Results obtained by extracting IL of 0–0.01 ppm insecticide solution with 20mL of solvent. Determination made by gas chromatography.
550
Insecticides and herbicides
Fig. 9.10 Screening technique described by Askew et al. [53].
Phosphorus-containing insecticides
551
Fig. 9.11 Comparison of gas chromatogram obtained with (i) a phosphorus detector and (ii) an electron-capture detector: (a) an extract equivalent to 1L of Thames water and (b) the same extract fortified with pesticides. Peaks correspond to 5ng of each compound: 1, Demeton-Smethyl; 2, Dimethoate; 3, Pyrimithate; 4, Parathion; 5, Chlorfenvinphos. (Reprinted with permission from Askew et al. [53]. Copyright (1969) Royal Society of Chemistry.)
with various organophosphorus insecticides. Figure 9.11 gives a comparison of the responses of an electron-capture detector and a phosphorus detector (flame thermionic); an Apiezon L column was used for the separation. Note the enhanced response for the chlorinecontaining insecticides, Chlorfenvinphos, on electron capture and the poor response for Demeton-S-methyl. The variation in the relative response of the electron-capture detector for organophosphorus compounds is a disadvantage for general screening. The interpretation of gas chromatograms obtained when using a phosphorus-specific detector of the Hartmann type must be made with caution. A chromatographic peak may often be caused by the presence of relatively large amounts of compounds that do not contain phosphorus; volatile nitrogencontaining compounds, in
552
Insecticides and herbicides
particular, can be detected at the microgram level. Additional confirmation of a characterization needs to be made with another technique, such as thin-layer chromatography. The pesticides amenable to gas chromatography can be separated on a variety of stationary phases; the separating times shown in Table 9.18 are based on columns that are commonly used for separating organochlorine insecticides. The retention times may vary greatly, and it is necessary to use either temperature programming or isothermal runs at more than one temperature for successful screening. The technique can be rendered quantitative down to subnanogram levels, and the comparison of relative times on two or more columns enables a tentative pesticide characterization to be made. The analytical scheme described by Askew et al. [53] is essentially a qualitative procedure but can be readily rendered quantitative. Those pesticides giving gas chromatographic peaks can be quantitatively determined by reference to a standard peak height– concentration curve. Blanchet [235] used hexane to extract organophosphorus insecticides from pond and marsh water prior to flame photometric gas chromatographic analysis. He also studied sample preservation techniques. Sixteen different organophosphorus insecticides were included in this study. Although Blanchet [235] found that hexane was not a good extractant for Amidithion, Vapona and Phosphamidon he found that these three compounds extracted with near 100% recovery into chloroform. Chloroform gave good results as an extractant for most of the other insecticides mentioned above, Daughton et al. [236, 237] have described a procedure for isolating and determining, in large volumes of aqueous media, ionic diethyl phosphate, diethyl thiophosphate, dimethyl phosphate and dimethyl thiophosphate suitable for application in environmental monitoring. Procedures for eliminating interference due to inorganic phosphate are also discussed. In this approach the aqueous sample containing ionic dialkyl phosphates and thiophosphates are passed down a column of Amberlite XAD-4 resin. Recoveries for diethylphosphoric acid and diethylthiophosphoric acid at 0.01–0.1ppm in 500–400mL of aqueous media were 100 and 85%, respectively; recoveries for dimethylphosphoric acid and dimethylthiophosphoric acid at 0.1ppm in 500mL of aqueous media were 50 and 97% respectively. Following a clean-up procedure the effluents are gas chromatographed using a gas chromatograph equipped with a phosphorus thermionic detector and a glass column (1.8m × 2mm i.d.), packed with equal parts of 15% QF-1 and 10% DC-200 on Gas-Chrom Q (80–100 mesh) at 140°C; injector, 200°C; detector, 250°C; nitrogen, 22mL min-1; air 23mL min-1; hydrogen, 55mL min-1.
Table 9.18 Usage and chromatographic results of organophosphorus insecticides
a
Notes A = agricultural, V = veterinary, F = food storage, H = horticultural. b The values shown are retention times relative to that of Parathion = 100. The columns used contained the following stationary phases (i) Apiezon L, 2% and Epikote 1001, 0.2%; (ii) SE30, 4% and Epikote 1001, 0.4%; and (iii) XE 60, 2% and Epikote 1001, 0.2%, coated on acid-washed, dimethyl dichlorosilane-treated 80–100 mesh Chromosorb G. All columns were 150cm in length with 0.3dm o.d. Retention times were determined at 195°C except where marked * = 220°C and † = 150°C. The retention times of Parathion on the three columns were (i) 220°C, 1.70min; 195°C, 4.0min; and 150°C, 19min; (ii) 220°C, 1.80min; 195°C, 4.50min; and 150°C 22min; and (iii) 220°C, 1.60min; 195°C, 4.25min; and 150°C, 21 min. ND denotes not detected. c The values shown are the R1 values in the solvent (i) hexane–acetone (5 + 1), (ii) chloroform–acetone (9+1) and (iii) chloroform–acetic acid (9 + 1). ‡ Denotes streaking. d The values shown are the elution volumes relative to that of Parathion = 100 when eluting the ethanol from a Sephadex LH 20 column.
Source: Reprinted with permission from Askew et al. [53]. Copyright (1969) Royal Society of Chemistry.
Table 9.18 Continued
Phosphorus-containing insecticides
555
Addition of chloroform to water samples containing traces of pesticides, as reported by Bourne [238] is not only a technique with numerous and tedious steps in the extraction procedure, but recovery percentages are lower for natural water samples. Moreover, traces of moisture in the extracts increase the degradation of the organophosphorus insecticides markedly and the recovery percentage decreases rapidly. The use of Amberlite XAD-2 resins as reported by Mallet and co-workers [239] is a breakthrough in the search for an adequate technique for the preservation of aqueous samples containing traces of organophosphorus insecticides. Puijker et al. [240] separated organic compounds containing phosphorus and sulphur from water on to XAD resin, then reduced the compounds with hydrogen at 1100°C. The resulting phosphine and hydrogen sulphide were separated on a chromatographic column, and detected at 526 and 384nm respectively, in a flame photometer. The detection limits for phosphorus and sulphur were 0.1ng and 1ng respectively. Cassita and Mallet [241] carried out a simultaneous d e t e r m i n a t i o n o f F e n i t ro t h i o n , A m i n o c a r b a n d t h e i r corresponding phenolic hydrolysis products in water. Phenols w e re d e r i v a t i z e d t o t h e c o r re s p o n d i n g e s t e r s w i t h a c e t i c anhydride directly in water. Extraction was with methylene chloride and simultaneous analysis achieved by gas–liquid chromatography using a nitrogen–phosphorus selective detector. The limit of detection was 0.1ug L -1 . Suffet and Faust [231] applied the p-value approach to the liquid– liquid extraction of Diazinon, Parathion, Malathion and Fenthion and their oxygen analogues and hydrolysis products from water samples prior to their analyses by gas chromatography on Reoplex-400 with electron-capture and flame ionization detection. Bargnoux et al. [232] carried out a comparative chromatographic study, utilizing thin-layer and gas chromatography with phosphorus and sulphur dual detectors for the detection of Parathion and Malathion residues in water. They discuss the application of two low temperature methods, lyophilization and cryoconcentration, for the recovery of these insecticides from water. The use of lyophilization and cryoconcentration represents an improvement in the analysis of some particularly labile phosphorus and sulphur pesticides, over previous methods by significantly reducing the risks of degradation in aqueous solution. Moreover, lyophilization offers two other advantages, by allowing on the one hand the simultaneous analysis of many water samples irrespective of their mineralization, and on the
Table 9.19 Recoveries of 38 organophosphorus pesticidal compounds
Source: Reprinted with permission from Thompson et al. [18]. Copyright (1977) Springer-Verlag, New York.
Phosphorus-containing insecticides
557
other, optimum conservation either in the freezing state before their treatment or after lyophilization step. Thompson et al. [18] have developed a multiclass, multiresidue gas chromatographic method for the determination of insecticides (organophosphorus, organochlorine, carbamate types) and herbicides in water samples. The compounds are extracted from water with methylene chloride, and the extract is concentrated by an evaporative technique utilizing reduced pressure and low temperature. Compounds are segregated into groups using a column of partially deactivated silica gel and sequential elution with four different solvent systems. Carbamate residues, converted to their 2,4-dinitrophenyl ether derivatives, are gas chromatographed via electron-capture detection as the parent compounds of the organohalogen compounds. Organophosphorus compounds are determined by gas chromatography using a flame photometric detector. Recovery studies were conducted on 42 halogenated compounds, 38 organophosphorus compounds (Table 9.19) and seven carbamates. Fifty-eight of 87 compounds tested produced recoveries in excess of 80%; another 13 compounds yielded recoveries exceeding 60%, while the remaining 16 compounds yielded recovered below 60%. The method is discussed in more detail in Section 9.1.1.1. Typical chromatograms of the flame photometric detection of the organophosphorus compounds are shown in Fig. 9.12. Thompson et al. [18] emphasize that even the use of two dissimilar gas chromatographic columns does not ensure irrefutable compound identification. For example, if the retention characteristics of a given peak obtained from two dissimilar columns suggest the possibility of the presence of a compound which appears wholly out of place in a specific sample, further confirmation is clearly indicated by such techniques as specific detectors, coulometry, p values, or gas chromatography–mass spectrometry or thin-layer chromatography. Gas chromatography with nitrogen–phosphorus or mass spectrometric detection has been used to determine organophosphorus pesticides in water samples [465]. Gas chromatography has been used [233] to determine the following organophosphorus insecticides at the microgram per litre level in water and waste water samples: Azinphos-methyl, Demeton-O, Demeton-S, Diazinon, Disulfoton, Malathion, Parathion-methyl and Parathionethyl. This method is claimed to offer several analytical alternatives, dependent on the analyst’s assessment of the nature and extent of interferences and the complexity of the pesticide mixtures found. Specifically, the procedure uses a mixture of 15% v/v methylene chloride in h e x a n e t o e x t r a c t o rg a n o p h o s p h o r u s i n s e c t i c i d e s f ro m t h e
558
Insecticides and herbicides
Fig. 9.12 Four organophosphorus compounds eluted in fraction II. Gas chromatographic column, 1.5% OV-171, 95% OV-210. (Reprinted with permission from Thompson et al. [18]. Copyright (1977) Springer Verlag, New York.)
aqueous sample. The method provides, through use of column chromatography and liquid–liquid partition methods for the elimination of non-pesticide interference and the preseparation of pesticide mixtures. Identification is made by selective gas chromatographic separation and may be corroborated through the use of two or more unlike columns. Detection and measurement are best accomplished by flame photometric gas chromatography using a phosphorus-specific filter. The electron-capture detector, though non-specific, may also be used for those compounds to which it responds. Confirmation of the identity of the compounds should be made by gas chromatography-mass spectrometry when a new or undefined sample type is being analysed and the concentration is adequate for such determination. Compounds such as organochlorine insecticides, PCBs and phthalate esters interfere with the analysis of organophosphorus insecticides by electron-capture gas chromatography. When encountered, these interferences are overcome by the use of the phosphorus-specific flame photometric detector. Elemental sulphur will interfere with the determination of organophosphorus insecticides by flame photometric and electron-capture gas
Phosphorus-containing insecticides
559
Fig. 9.13 Column packing: 1.5% OV-17 + 1.95% QF-1. Carrier gas: nitrogen at 70ml min-1. Column temperature, 215°C. Detector, flame photometric (phosphorus).
chromatography. The elimination of elemental sulphur as an interference is discussed in detail. A typical gas chromatogram obtained by this procedure are shown in Fig. 9.13. Retention data for various gas chromatographic column packings are reproduced in Table 9.20. Ve r w e i j e t a l . [ 2 3 4 ] h a v e d e s c r i b e d a p ro c e d u re f o r t h e determination of PH -containing insecticides in surface water. In 3 this procedure the insecticide is hydrolysed to methylphosphonic acid, and the acid is concentrated by anion exchange and converted to the dimethyl ester. After clean-up on a microsilica gel column the ester is analysed by gas chromatography using a thermionic phosphorus-specific detector. Detection limit is 1nmol L -1. The gas chromatographic analysis was carried out on a Becker gas chromatograph, type 409, equipped with a thermionic detector, type 712. The coiled glass column (2m × 1.5mm i.d.) was packed with Chromosorb W AW/DMCS 80–100 mesh coated with Triton X-305 (25% w/w after sieving in the particle range 149–177µm. The column, injector and detector were maintained at 150, 100 and 200°C respectively. The carrier glass flow rate was 40mL min-1 of nitrogen. Because of the use of a splitter at the end of the column (ratio 3:1)
560
Insecticides and herbicides
Table 9.20 Retention times of some organophosphorus insecticides relative to Parathion
Notes a All columns glass, 180cm × 4mm i.d., solid support Gas-Chrom Q, 100–120 mesh. b May substitute OV-210 for QF-1. c Anomalous, multipeak response often encountered.
only 30mL of nitrogen per minute reached the thermionic detector. The remaining part was led to a flame ionization detector. Between 0.2 and 1.2µg L-1 PH was found by this method in rivers 3 flowing through Holland. The source of this contamination has not been identified. Mecarphon, the only PH -containing insecticide is 3 supposedly no longer produced. Detergents and flame retardants are other possibilities. To prove that a peak ascribed to dimethyl methylphosphonate is not due to the presence of a non-phosphorus compound in relatively high concentrations, the thermionic detector was used in combination with a flame ionization detector. In the case of a non-phosphorus compound the last-mentioned detector will give a relatively high response. The identity of the compound giving rise to this peak was confirmed by simultaneous injection of a reference sample of dimethyl methylphosphonate as well as by mass fragmentography. 9.3.1.4 Thin-layer chromatography Various thin-layer chromatographic systems have been described for the identification and determination of organophosphorus insecticides in natural waters (Table 9.21). Sherma and Boymel [242] determined Abate in natural water samples by densitometry on preadsorbent reversed-phase thin-layer plates.
Table 9.21 Thin-layer chromatography of organophosphorus insecticides
Table 9.21 Continued
Phosphorus-containing insecticides
563
9.3.1.5 High-performance liquid chromatography Liquid chromatography has been used to determine thermally labile organophosphorus pesticides in water [466, 467]. Lacorte and Barcelo [468] have described a procedure for the determination of nanogram per litre levels of organophosphorus pesticides in ground waters based on automated on-line liquid–solid extraction followed by liquid chromatography. The detection used on the liquid chromatograph is an atmospheric pressure chemical ionization mass spectrometer using negative- and positive-ion modes of operation. The technique was used to determine several organophosphorus pesticides, (E)- and (Z)-Mevinphos, Dichlorvos, Azinphos-methyl, Azinphosethyl, Parathion-methyl, Parathion-ethyl, Malathion, Fenitrothion, Fenthion, Chlorfenvinphos and Diazinon, in ground water. This method required 100mL of water, and it was combined with a prior automated on-line liquid–solid extraction step using an OSP–1 autosampler containing C cartridges. The limit of quantitation 18 varied between 5 and 37ng L -1 in positive-ion (PI) mode. Under negative-ion (NI) mode of operation, only the Parathion group (both Parathions and Fenitrothion) had a better sensitivity as compared to that in PI mode, with a limit of quantitation of 5–515ng L-1, whereas the rest of the pesticides had two to four times higher limits of quantitation as compared to those in PI mode. Selected ion monitoring of the group-specific fragment of the organophosphorus pesticides, e.g., [(CH O) PO ] - or the [M + H] + ions, under NI or PI mode, 3 2 2 respectively, was used. Sample cone voltage varied from 10 to 30V, good fragmentation was obtained for most of the studied analytes. The system was used for the certification of a ground water sample spiked at the nanogram per litre level with organophosphorus pesticides provided by Aquachek. Farren et al. [243] have discussed a method of continuous flow extraction of organic phosphorus insecticides coupled on-line with highperformance liquid chromatography. Simon [244] has described a high-performance liquid chromatography based on an assay method for the determination of down to 1µg L-1 Paraquat and Diquat in well water. Paraquat was detected by UV absorption at 257nm and Diquat at 310nm. 9.3.1.6 Reverse-phase adsorption liquid chromatography Otsuki and Takaku [245] have described a method for the determination of Abate down to 5–150µg using reversed-phase adsorption liquid chromatography. Only 1mL of sample is required for this determination.
564
Insecticides and herbicides
9.3.1.7 Raman spectroscopy Thibeau et al. [246] studied the feasibility of applying preresonance Raman spectroscopy for detection and identification of hazardous chemicals present in water in trace amounts. Several pesticides and fungicides including Parathion-methyl, all based on the nitrobenzene structure were selected on account of their partial solubility in water. Details of the absorption band maxima, minimal detectable concentrations and frequencies of observed Raman bands for several typical compounds are presented. 9.3.1.8 Polarography Brøstad et al. [255] have described a polarographic method for the determination of glyphosphate residues in natural waters as their Nnitroso derivatives. 9.3.1.9 Enzymic assay methods Gauson et al. [256] have described a method using a source of cholinesterase enzyme impregnated on paper for the rapid and simple detection of organophosphorus inhibitors in water in concentrations at or about the level of acceptability for drinking contaminated water. The papers are immersed in a suspect water and after a suitable incubation time, a chromogenic substrate is added to the paper. The appearance of a blue colour, different from that of the substrate is evidence that the enzyme is active and that the water does not contain sufficient anticholinesteratic material to inhibit the enzyme. Conversely, no change in the colour is evidence that an anticholinesteratic material is present in a concentration equal to or greater than that acceptable for ingestion. These workers give experimental data on studies with the organophosphorus esters isopropyl methylphosphonofluoridate (Sarin), O-ethyl-S-(2-diisopropyamino)ethyl methylphosphonothioate, and O-(4-nitrophenyl phosphorothioate) (Parathion). Karlhuber and Eberle [257] have discussed the determination of organophosphorus and carbamate insecticides by cholinesterase inhibition. Goodson et al. [258] used an immobilized cholinesterase product for the detection of organophosphorus and carbamate insecticides in water. In this method, pads in which horse-serum cholinesterase is entrapped in starch gel on the surface of open-pore polyurethane foam [259] were used to detect enzyme inhibitors in air or water. Pads with improved enzyme activity and better resistance to loss of such activity are
Phosphorus-containing insecticides
565
prepared by adsorption of cholinesterase on aluminium hydroxide gel during precipitation of aluminium hydroxide from a solution of aluminium chloride followed by suspension of the gel enzyme mixture in a slurry of starch for application to the polyurethane foam. Pads so prepared retain their activity during the monitoring of water for certain organophosphorus and carbamate insecticides. The US Environmental Protection Agency [260] has also reported on a rapid detection system based on the use of immobilized cholinesterase for the detection of organophosphorus and carbamate insecticides in water. 9.3.1.10 Miscellaneous Addition of chloroform to water samples containing traces of pesticides, as reported by Bourne [238], is not only a technique with numerous and tedious steps in the extraction procedure, but recovery percentages are lower than for natural water samples. Moreover, traces of moisture in the extracts increase the degradation of the organophosphorus insecticides markedly and the recovery percentage decreases rapidly. Sharma et al. [261] described a specific spot test to detect the presence of Malathion in acidic, basic and saline waters. Malathion is concentrated from the sample on activated carbon, extracted with ethanol, and hydrolysed with potassium hydroxide to form potassium fumarate which gives a red colour when heated with acetic anhydride. Goosen and Kloosterboer [262] have described a method for the determination of organophosphorus-derived orthophosphates in natural and waste water after photochemical ultraviolet decomposition and acid hydrolysis. The use of Amberlite XAD-2 resins as reported by Mallet and coworkers [239] is a breakthrough in the search for an adequate technique for the preservation of aqueous samples containing traces of organophosphorus insecticides. Vogler [263] has described a method for the separate determination of orthophosphoric acid esters (‘COP-phosphate’) and condensed phosphates (‘POP-phosphate’) in natural water samples. The sample (100mL) of filtered natural water is treated with 30% aqueous hydrogen peroxide (0.2mL) and exposed to UV radiation at below 23°C under conditions of maximum irradiation intensity and minimum time. Excess amounts of hydrogen peroxide and orthophosphates are determined by standard procedures. The result obtained is a measure of COPphosphate. The other half of the sample is treated in a polyethylene flask with 10N sulphuric acid (0.4mL) on a steam bath for 4h. Excess hydrogen peroxide is then removed with potassium permanganate solution and the orthophosphate is again determined, with a waiting
566
Insecticides and herbicides
time of 20min after the addition of reagent. The value obtained is for total dissolved phosphate; POP-phosphate is obtained by subtraction. Russell Everett and Rechnitz [469] have described a mediated electrocatalytic method incorporating a tyrosinase-based oxygen biosensor for the determination of organophosphorus insecticides in water samples. The tyrosinase-based oxygen sensor is constructed in such a way that enzymatic oxygen consumption is monitored electrochemically using the mediator 1,2-naphthaquinone-4-sulphonate. This bioelectrocatalytic system allows electrochemical initiation and termination of the enzymatic reaction. Amperometric and coulometric techniques were used by these workers to study inhibiting effects in pesticide determinations. This inhibition appears to be fully reversible, with full catalytic activity returning after the removal of pesticides. Detection limits of 5µM were achieved for Diazinon and 75µM for Dichlorvos. Enzymatic inhibition occurs at approximately 1000 and 50µM respectively. Height flow pneumatically assisted electrospray under positive-ion mode permitted program levels of organophosphorus pesticides, e.g., Parathion-methyl, Parathion-ethyl, Fenitrothion and Fenitrooxon, to be determined in water samples [470, 471]. Ting-Long et al. [487] have described a selective determination of Dimethoate in water using ion-mobility spectrometry and mixed alternate reagent ions. By doping ammonium, acetone, ethanol, diethyl ether and methanol into the carrier gas, it was shown that the best selective detection was obtained by the addition of ammonium ion, which had the highest proton affinity of all the above reagents. Three mixed alternate reagents to improve selectivity, viz. ammonium ion– acetone, ammonium ion–diethyl ether and acetone– diethyl ether, were studied. It was shown that single-solvent dopants were advantageous. The best resolution and highest abundance of the Dimethoate peak was observed in its ion mobility spectrum after the addition of the mixed dopant ammonium ion–acetone. Ammonium ion at 1g L-1 plus acetone at 1g L-1 gave the strongest response for Dimethoate. 9.3.1.11 Preconcentration Bargnoux et al. [232] used preconcentration at low temperatures as a means of extracting organophosphorus insecticides from water, prior to gas chromatography. Volpe and Mallet [264] developed a method for determining down to 0.5ng of Fenitrothion and five Fenitrothion derivatives in water by adsorption on XAD-4 and XAD-7 resins, followed by solvent elution and gas–liquid chromatography of the extract.
Phosphorus-containing insecticides
567
9.3.1.12 Individual organophosphorus insecticides Much work has been published which discusses the determination of individual organophosphorus insecticides. Whilst, as has already been discussed, in general it is advisable to adopt multicompound approaches when analysing for this class of compounds or mixtures of these with organochlorine and carbamate insecticides and herbicides and PCBs, there are circumstances when a simple direct method for a particular organophosphorus insecticide might be useful. Such methods are discussed in this section.
Parathion (O-(4-nitrophenyl phosphorothioate)), Parathion-methyl (dimethyl-p-nitrophenyl phosphorothioate), Parathion-ethyl (diethyl-p-nitrophenyl phosphorothioate) Kawahara [25] described a procedure for the determination of phosphorothioate insecticides including Parathion and Parathionmethyl. It consists of solvent extraction, clean-up by thin-layer chromatography on silica gel G (0.25mm layer), and identification by gas chromatography on an aluminium column (1.2m × 0.6cm o.d.) packed with equal portions of acid-washed Chromosorb P supporting 5% of DC 200 silicone oil, and unwashed Chromosorb W supporting 5% of Dow-11 silicone: the column is operated at 180°C, with argon-methane (9:1) as carrier gas (120mL min-1) and electron-capture detection. If the sample volume is sufficient, identification can be confirmed by infrared spectrometry. This method was used to follow accidental contamination of river water by pesticides. Other workers [265–267] have used gas chromatography to determine Parathion in water samples. Moye [268] has discussed the application of liquid chromatography to the determination of Parathion-ethyl. Paschal et al. [269] have discussed the determination of Parathionethyl and Parathion-methyl in run-off water using high-performance liquid chromatography. The organic compounds are concentrated on an XAD-2 resin before analysis by reversed-phase, high-performance liquid chromatography. Detection limits were found to be approximately 2–3mg L1 . These workers examined the possible interferences to the method from other agricultural chemicals and organic compounds commonly occurring in water. This method is based on the use of Rohm and Haas XAD macroreticular resins [163]. Organics in water can be sorbed on a small column of resin, and the sorbed organics then eluted by diethyl ether. After evaporation of the eluate, the concentrated organics can be determined by chromatography. In addition to the obvious benefit of 100–1000-fold concentration, this method offers the possibility of on-site sampling.
568
Insecticides and herbicides
Venkataraman and Sathyamurthy [201] developed a simple and direct spectrophotometric method for the determination of Parathion in water in which a benzene extract of Parathion is hydrolysed and is reduced with zinc dust in acid solution. The resulting amino derivative is diazotized with sodium nitrite and hydrochloric acid. The diazo compound is coupled with naphthylethylenediamine hydrochloride and a magenta dye is produced, which is evaluated spectrophotometrically. It is unlikely that this method is specific for Parathion. Malathion Bargnoux et al. [232] carried out a comparative chromatographic study, utilizing thin-layer and gas chromatography with phosphorus and sulphur dual detectors for the detection of Parathion and Malathion residues in water. They discuss the application of two low temperature methods, lyophilization and cryoconcentration for the recovery of these insecticides from water. The use of lyophilization and cryoconcentration represents an improvement in the analysis of some particularly labile phosphorus and sulphur pesticides over previous methods by significantly reducing the risks of degradation in aqueous solution. Moreover, lyophilization offers two other advantages, by allowing on the one hand the simultaneous analysis of many water samples irrespective of their mineralization, and on the other, optimum conservation either in the freezing state before their treatment or after the lyophilization step. Wolfe et al. [270] studied the kinetics of the chemical degradation of Malathion in water. They investigated potential chemical and photochemical pathways for the degradation of Malathion. The results of acid-catalysed degradation and oxidation indicate that these pathways would be too slow to be of significance in the aquatic environment, but photolysis could be important under acid conditions. Malathion was much more susceptible to alkaline degradation, and underwent two competing reactions at 27°C. The persistence of the Malathion monoacids formed was also studied, and they proved to be 18 times more stable than Malathion under the same conditions. Dursban (O,O-diethyl-O-(3,5,6-trichloro-2-pyridyl phosphorothioate) A gas chromatographic procedure using electron-capture detection has been described for the determination of Dursban in water and silt (Rice and Dishberger [271]). In this method, water samples are extracted with dichloromethane, the extract is evaporated, and a solution of the residue is cleaned up on a column of silicic acid, Dursban being eluted
Phosphorus-containing insecticides
569
with hexane. The eluate is evaporated to dryness under reduced pressure, and a solution of the residue in hexane is subjected to gas chromatography on a glass column (4ft × 0.25in o.d.), packed with 5% of SE-30 on Anakrom ABS (80–90 mesh) operated at 200°C (or 215°C with methane–argon (1:19) (60mL min-1) as carrier gas, or, 5% SF-96 on Chromosorb W (80–100 mesh), operated at 215°C with nitrogen (90mL min-1) as carrier gas. Dried silt samples, finely powdered, are blended with dichloromethane and Celite, the filtered extract is evaporated to dryness and a solution of the residue in hexane is extracted with acetonitrile. The concentrated solution plus added hexane is evaporated to dryness under reduced pressure, and Dursban in a solution of the residue in hexane is subjected to clean-up and gas chromatographed as for water samples. Down to 10-4ppm of Dursban in water and down to 5 × 10-3ppm in silt could be determined; average recoveries from water and silt were 92% and 83% respectively. Deutsch et al. [272] determined Dursban in water, mud, vegetation, fish, ducks, insects and Crustacea. After a preliminary clean-up the extract is chromatographed on a column packed with 3% Carbowax 20M on Gas-Chrom (60–80 mesh), which gives excellent separation of Dursban from other organophosphorus insecticides. Both thermionic and flame photometric detectors are satisfactory. Recoveries range from 75 to 105% depending on the nature of the sample. This procedure will detect as little as 0.5ng of Dursban, corresponding to a level of 0.01ppm, in a 10g sample. Abate (O,O,O’,O’-tetramethyl-O,O’-(thiodi-p-phenylene)-diphosphorothioate) Dale and Miles [273] showed that Abate and its sulphoxide were well separated from various other organophosphorus insecticides on a column of XE-60 on silanized Chromosorb W at 240°C, using nitrogen as carrier gas and a flame photometric and electron-capture detector. Shafik [275] has described a method for the determination of this insecticide in water. Abate is first converted to its hydrolysis product, 4,4'-thiodiphenol, which is then silated by reaction with chlorotrimethylsilane and hexamethyldisilazane. Separation was achieved on an aluminium column (4ft × 0.25in o.d. (1.2m × 6mm)) containing 2.5% E-301 or 0.25% Epon 1001 on Chromosorb W (80–100 mesh) operated at 190°C with nitrogen as carrier gas (100mL min-1) and a flame photometric detector equipped with a sulphur filter. Miller and Funes [274] used alkali flame gas chromatography to determine Abate. Separation was achieved on a column packed with 2.5% of E-301 plus 0.25% of Epon 1001 on Gas-Chrom W (AW-DMCS)HO, the column and detector being operated at 235°C. The column was conditioned by injecting 20ng standards for up to 200ng Abate. The recovery of 0.01 to
570
Insecticides and herbicides
1ppm of added Abate to water was 97%, after extraction by the method of Dale and Miles [273]. Thin-layer chromatography has also been applied to the determination of Abate residues in water [276]. The sample of surface water or sewage, acidified with sulphuric acid, was extracted with chloroform. The extract was evaporated under nitrogen at 60–70°C and the residue was dissolved in acetone; 15µl of this solution was applied to a layer (0.25mm) of silica gel G, previously activated for 1h at 90°C, together with Abate standards. The plate was developed for 1h in hexane– acetone (10:1), air dried for 1min, exposed to bromine vapour for 1min, exposed to air for 3min, and sprayed with 1% N,N-dimethyl-p(phenylazo)aniline (CI solvent yellow 2) solution in 95% ispropyl alcohol. Abate gave distinct red spots (R 0.11 ± 0.01) on a bright yellow f background. Spot areas were related to concentration by a calibration graph. Down to 9µg of Abate could be readily determined. Otsuki and Takaku [245] have described a method for the determination of Abate down to 5–150µg using reversed-phase adsorption liquid chromatography. Only 1mL of sample is required for this determination. S-Methyl Fenitrothion (O,S-dimethyl-O-(3-methyl-4-nitrophenyl)phosphorothiolate) Zitko and Cunningham [279] investigated the hydrolysis rate of this compound and prepared ultraviolet, infrared and mass spectra. Fenitrothion Various workers [236, 280–282, 277, 278] have used macroreticular resin XAD-2 to recover Fenitrothion from river water samples and compared this method with solvent extraction techniques. Mallet et al. [280] used an automated gas chromatographic system which consisted of a gas chromatograph mounted with an automatic sample interfaced to an integrator. A Melpar flame photometric detector (phosphorus mode) was connected with the flame gas inlets in the reverse configuration to prevent solvent flame-out. The detector was maintained at 185°C and flame gases were optimized with flow rates (mL min-1) as follows: hydrogen, 80; oxygen, 10; air, 20. A 2.8m × 4.0mm i.d. U-shaped glass column packed with 4% (w/w) OV-101 and 6% (w/ w) OV-210 on Chromosorb W AW DCMS, 80–100 mesh was used. Nitrogen was used as carrier gas at a flow rate of 70mL min-1. A column temperature of 195°C sufficiently resolved the Fenitrooxon from its parent compound. The injection port temperature was set at 225°C. The water sample was passed through an XAD-2 column, which was
Phosphorus-containing insecticides
571
subsequently eluted with ethyl acetate. The ethyl acetate extract was examined by gas chromatography. Methods [283, 284] to determine Fenitrothion in water using gas– liquid chromatography with a flame photometric detector have been reported. An in situ fluorimetric method [285] to detect simultaneously Fenitrothion, Fenitrooxon, Aminofenitrothion and nitrocresol on a thinlayer chromatogram has been developed. Fenitrothion, Fenitrooxon and Aminofenitrothion have been analysed simultaneously by gas chromatography using an SE-30 plus QF-1 column and a flame photometric detector in the phosphorus mode [286]. Coumaphos (Coral) and Bayrusil Mallet and Brun [287] have described an in situ spectrofluorometric method for the analysis of these substances in lake water. This procedure does not necessitate a preliminary cleaning up of samples, and interfering coextractives which fluoresce only in solution are also avoided. An advantage over other in situ fluorimetric techniques is that fluorogenic spray reagents are not used and the fluorescence produced is selective. In earlier papers Brun and Mallet [288] had shown that Coral and Bayrusil could be made to fluoresce on silica gel layers simply by heating the chromatogram at a specific temperature for a definite period of time. Recoveries obtained by this procedure ranged from 86 to 108% in the concentration range 0.1 to 10µg L-1 water for Coral and from 86 to 94% in the concentration range 1 to 10µg L-1 water for Bayrusil. 9.3.2 Sea water 9.3.2.1 Spectrophotometric method Weber [289] has described a kinetic method for studying the degradation of Parathion in sea water. Weber observed two pathways whereby Parathion is hydrolysed. The first reaction proceeds via dearylation with loss of p-nitrophenol: (C2H5O)2 - PS - OC6H4 - NO2 + H2O = (C2H5O)2 - PS - OH + HOC6H4NO2 Additionally, they observed a second pathway, hydrolysis through dealkylation leading to a secondary ester of phosphoric acid which still contains the p-nitrophenyl moiety, i.e. de-ethyl Parathion (O-ethyl-O-pnitrophenyl-monothiophosphoric acid):
572
Insecticides and herbicides
(C H O) - PS - OC H4NO + H O 2 5 2 6 2 2 = NO - C H O - PS(OC H )OH + C H OH 2
6
4
2
5
2
5
Aliquots of ethanolic solution of Parathion were separated into undecomposed insecticides and decomposed insecticide products. Among the products free p-nitrophenol, chemically bound in acidic phosphorus compounds and in non-hydrolyzed neutral phosphorus compounds, was detected in the same way after saponification. Saponification after removal of ether without separation of neutral and acidic compounds yielded total p-nitrophenyl equivalents. 9.3.3 Potable waters 9.3.3.1 Gas chromatography Cranmer and Peoples [290, 291] have also described a method for determining anticholinesterase pesticides in potable water. Compounds that are direct inhibitors can be differentiated from those that must be activated with bromine before they possess anticholinesterase activity. The method involves the incubation of various amounts of non-treated and bromine treated sample with acetylcholinesterase for 1h at 37°C, addition of substrate (3,3-dimethylbutyl acetate) and, after a further 30min at 37°C, stopping the reaction by adding formic acid and then extracting with carbon disulphide. The reaction product (3,3-dimethylbutanol) in an aliquot of the extract is determined by gas chromatography and the inhibited samples. 9.3.4 Waste waters 9.3.4.1 Gas chromatography Gas chromatography has been used [292] to determine the following organophosphorus insecticides at the microgram per litre level in water and waste water samples: Azinphos-methyl, Demeton-O, Demeton-S, Diazinon, Disolfoton, Malathion, Parathion-methyl and Parathionethyl. This method is claimed to offer several analytical alternatives, dependent on the analysts’ assessment of the nature and extent of interferences and the complexity of the pesticide mixtures found. Specifically, the procedure uses a mixture of 15% v/v methylene chloride in hexane to extract organophosphorus insecticides from the aqueous sample. The method provides, through use of column chromatography and liquid–liquid partition, methods for the
Phosphorus-containing insecticides
573
elimination of non-pesticide interference and the preseparation of pesticide mixtures. Identification is made by selective gas chromatographic separations and may be corroborated through the use of two or more unlike columns. Detection and measurement are best accomplished by flame photometric gas chromatography using a phosphorus-specific filter. The electron-capture detector, though nonspecific, may also be used for those compounds to which it responds. Confirmation of the identity of the compounds should be made by gas chromatography– mass spectrometry when a new or undefined sample type is being analysed and the concentration is adequate for such determination. Detailed instructions are given for clean-up of reagents, solvents and glassware to avoid the occurrence of discrete artefacts and/or elevated baselines. Sample clean-up procedures are generally required and may result in the loss of certain organophosphorus insecticides. Therefore, great care should be exercised in the selection and use of methods for eliminating or minimizing interferences. Compounds such as organochlorine insecticides, polychlorinated biphenyls and phthalate esters interfere with the analysis of organophosphorus insecticides by electron-capture gas chromatography. When encountered, these interferences are overcome by the use of the phosphorus-specific flame photometric detector. Elemental sulphur will interfere with the determination of organophosphorus insecticides by flame photometric and electroncapture gas chromatography. The elimination of elemental sulphur as an interference is discussed in detail. 9.3.4.2 Field desorption mass spectrometry Schulten and Sun [293] used field desorption mass spectrometry to identify organophosphorus insecticides in waste water. The characteristic behaviour of four groups of common organophosphorus pesticides (phosphate, phosphorothionates, phosphorothiolates and phosphorodithioates) were investigated. This technique is suitable for identification of traces of these compounds in waste waters and other environmental samples without the need for preliminary separation and purification. 9.3.4.3 High-performance liquid chromatography Liquid–liquid extraction or preconcentration adsorption resins have been combined with high-performance liquid chromatography to determine organophosphorus insecticides in waste waters [294].
574
Insecticides and herbicides
9.3.5 Sewage effluents 9.3.5.1 Gas chromatography McIntyre et al. [295] investigated methods for determining organophosphorus compounds in sewage sludge and made a survey of sewage sludges from 12 different UK sewage treatment works. Highest recoveries were achieved using the disperser method, together with an alumina column clean-up step, and analysis by flame photometric gas– liquid chromatography. The survey results indicate that organophosphorus compounds are not very common micropollutants of sewage sludges. 9.3.5.2 Thin-layer chromatography Thin-layer chromatography has also been applied to the determination of Abate residues in water [276]. The sample of surface water or sewage, acidified with sulphuric acid, was extracted with chloroform. The extract was evaporated under nitrogen at 60–70°C and the residue was dissolved in acetone. This solution was applied to a layer of activated silica gel G for 1h at 90°C together with Abate standards. The plate was developed for 1h in hexane–acetone (10:1), air dried for 1min, exposed to bromine vapour for 1min, exposed to air for 3min, and sprayed with 1% N,N-dimethyl-p(phenylazo)aniline (C1 solvent yellow 2) solution in 95% isopropyl alcohol. Abate gave distinct red spots (R 0.11 ± 0.01) on a bright yellow f background. Spot areas were related to concentration by a calibration graph. Down to 9µg of Abate could be readily determined. 9.4 Pyrethrin insecticides 9.4.1 Natural waters 9.4.1.1 Gas chromatography Down to 0.2µg pyrethrin can be determined in water by gas chromatography [296]. A hexane extract of the sample is washed with aqueous sodium chloride solution; then follows an elaborate working-up procedure, and it is gas chromatographed on a helical glass column (4ft × 0.25in (1.2m × 6mm)) packed with 5% of SE-30 on AW-DMCS Chromosorb W (60–80 mesh) and operated at 190°C with nitrogen (40mL min-1) as carrier gas and flame ionization detection. This permits the simultaneous determination of pyrethrin (linear response range 0.2– 2.2µg) and of the synergistic piperonyl butoxide (range 0.6–5.6µg) and N-
Carbamate insecticides and herbicides
575
(2-ethylhexyl)-Norborn-5-ene-2,3-dicarboximide (range 0.6–1.8µg). Recoveries averaged 93–94%. 9.5 Thiram (tetramethylthiruam) insecticides 9.5.1 Natural waters 9.5.1.1 Spectrophotometric method This substance can be determined in waters spectrophotometrically [297]. A chloroform extract of the sample is reacted with a methanolic suspension of cupric iodate and the extraction of the cream-yellow colour evaluated spectrophotometrically at 440nm. 9.5.1.2 Voltammetry Procopio et al. [298] have determined Thiram in natural waters at concentrations down to 0.03µg L-1 by a by a voltammetric procedure. 9.6 Carbamate insecticides and herbicides 9.6.1 Natural waters 9.6.1.1 Spectrophotometric methods To determine traces of Carbine (Barban (4-chlorobut-2-ynyl-3-chlorocarbanilate)) in water, Bosyakova et al. [299] distilled the sample with sulphuric acid, collecting the distillate. The residue was hyrolysed to mChloroaniline with dilute sodium hydroxide and the product steam distilled. m-Chloroaniline was determined spectrophotometrically following diazotization with nitrous acid and 2-naphthol. Handa and Dikshit [300] have described a Spectrophotometric method for the determination of Carbaryl in lake, stream and pond waters. This method is based on the use of vanillin as chromogenic reagent. Bendiocarb has been determined spectrophotometrically in fresh water as its complex with nitric acid [301]. Furadan (Carbofuran) has been determined spectrophotometrically as a reaction product with sulphuric acid and ceric sulphate in natural water [302]. Condo and Janauer [303] give details of a procedure for the indirect Spectrophotometric determination of traces of the pesticide Aldicarb in water. It involved decomposition of Aldicarb by passage through a
576
Insecticides and herbicides
strongly basic anion-exchange resin to produce an oximate which was adsorbed on the resin. Subsequent addition of sulphuric acid resulted in in situ formation and elution of hydroxylamine, which was used for quantitative reduction of trivalent iron to the ferrous state; this was then determined spectrophotometrically. The carbamate herbicide Propham has been determined by synchronous derivative spectrophotometry following fluorescamine fluorogenic labelling [304]. Propham was hydrolysed with 1M sodium hydroxide at 100°C for 40min. The liberated aniline reacted with fluorescamine in a potassium phthalate buffer. Fluorescamine labelling was followed by synchronous derivative spectrofluorimetry. Reaction rates, fluorescence phenomena and synchronous derivative parameters were investigated. Interference by naphthaleneacetic acid, colchicine, (2,4-dichlorophenoxy)-acetic acid, Flurecol and Dimethoate was examined. The minimal detectable quantity was estimated to 7.2µg L-1 Propham. The range of determination was 24–2000µg L-1. Various workers have described spectrophotometric methods for the determination of Carbaryl in natural waters [305–307]. 9.6.1.2 Gas chromatography Probably the most common of insecticides is Carbaryl. This substance has obtained wide acceptance due to its effectiveness and low mammalian toxicity. Crosby and Bowers [308] have described a method for determination of Carbaryl (Sevin) (1-naphthyl-N-methylcarbamate) in which the sample (0.5g) is heated under reflux for 1h with 2-chloro-a,a,a-5nitroluene (1mmol) or 4-chloro-a,a,a-trifluoro-3,3-dinitroluene (0.7mmol), acetone (20mL) and 0.1M Na B O (20mL) to convert its amine moiety 2 4 7 into an N-substituted nitro-(trifluoromethyl)aniline derivative. The derivative is subjected to gas chromatography on a stainless steel column (10ft × 0.125in o.d. (3m × 3mm)) packed with 3% SE-30 or 3% FFAP on HMDS treated Chromosorb G at a temperature from 150 to 250°C, with nitrogen (30mL min-1) as carrier gas and electron capture or flame ionization detection. Down to 50pg of the more volatile or 200pg of the less volatile derivatives can be determined by electron-capture detection. Methods also described are the determination of Molinate (Sethyl hexamethylenecarbamate) in water. Methods for analysing mixtures of carbamate insecticides have been described by various workers [18, 309]. Cohen et al. [309] used an electron capture gas chromatograph to determine Carbamate insecticides as their 2,4-dinitrophenyl derivatives in water and plant material. After isolation from the sample material by the appropriate extraction method, the carbamates were reacted with 1-fluoro-2,4-dinitrobenzene. The
Carbamate insecticides and herbicides
577
resulting 2,4-dinitrophenyl ethers were determined by subjecting their solution in hexane to gas chromatography on a glass column (140cm × 15mm) packed with 1% GE-XE 60 and 0.1% Epikote 1001 on AW-DMCS Chromosorb G (60–80 mesh) operated at 211°C; nitrogen was used as carrier gas. The recovery of carbamates added to water and to peas, lettuce and apples was in the range of 82–100%, except for Carbaryl (63%) and Butacarb (40–65%) added to water. The limits of determination of carbamates in water and plant materials were 0.005ppm and 0.1ppm respectively. Reeves and Woodham [311] have described a gas chromatographic method for the determination of Methomyl [464] (S-methyl-N-((methylcarbamoxyl)oxy)thioacetimidate) in water. The residues were extracted from water with dichloromethane, and the extracts were purified on a column of Florasil. The extracts were purified by a coagulation procedure with ammonium chloride–phosphoric acid. The purified and concentrated extracts were then analysed by gas chromatography on a glass column (6ft × 3/16in (1.8m × 5mm)) packed with 10% DC-200 on Chromosorb W HP (80–100 mesh) and operated at 140°C, with nitrogen as carrier gas (80mL min-1) and a 394nm S-interference filter. The limit of detection was 0.01ppm for water; the recovery was 75.1%. Westlake et al. [310] determined m-S-butylphenyl methyl(phenylthio)carbamate (RE11775) in water, by a gas chromatographic procedure. The sample is extracted with dichloromethane, chloroform or acetonitrile, followed by clean-up, if necessary, on a column of Florasil, silica gel or alumina. The purified residue is submitted to gas chromatography on either a stainless steel column (3ft × 0.25in (0.9m × 6mm)) packed with 5% OV-225 on Gas-Chrom Q (60–80 mesh) and operated at 242°C, with nitrogen as carrier and a flame photometric detector operated in the S mode, or on a glass column (3ft (0.9m) × 6mm o.d.) with identical packing and operated at 195°C, with hydrogen as carrier gas (100mL min-1) and an electrolytic conductivity nitrogen detector. Recoveries of added RE11775 from water were about 100% and from grass and lucerne about 80%. Down to 0.01ppm could be determined in water. Various workers [309, 312–316] have discussed gas chromatographic methods for the determination of Carbaryl in water samples. Lewis and Paris [316] determined Carbaryl and its hydrolysis product 1-naphthol by direct injection of a benzene extract on Gas-Chrom Q (80–100 mesh) and operated at 145°C with nitrogen (120mL min-1) as carrier gas. Considerable preconditioning of the column was necessary to obtain optimum sensitivity. Thompson et al. [18] have devised a multiresidue scheme of analyses based on silica gel column chromatography followed by gas chromatography for the analyses of mixtures of organochlorine, organophosphorus and carbamate types of insecticides. The relevant work
578
Insecticides and herbicides
on seven carbamate insecticides is discussed below. The mixture of the three types of compounds is fractionated into groups on a partially deactivated silica gel column with three sequential elutions. Final determinations were made by gas chromatography using the carboncapture detector for the halogenated compounds and derivatized carbamates, and the flame photometric detector for the organophosphorus compounds. Derivatization of the carbamate fraction is carried out with 1fluoro-2,4,-dinitrobenzene as follows. To the tubes containing the 0.1mL concentrates of the carbamate fractions is added 0.5mL of 1-fluoro-2,4dinitrobenzene (1% in acetone) and 5mL of sodium borate buffer solution (Na B O -10H O, 1M solution at pH0.4). The reagents are added to an 2 4 7 2 empty tube to serve as a reagent blank. The tubes are tightly capped and heated at 70°C for 1h in a water bath. The tubes are cooled to room temperature, and 5mL of hexane is added to each tube. The tubes are shaken vigorously for 3min, either manually or on a wrist action shaker. The layers are allowed to separate and 4mL of the hexane (upper) layer are transferred carefully to a tube and stoppered tightly. The results obtained in applying electron-capture gas chromatography to a mixture of six carbamate insecticides is shown in Fig. 9.14.
Fig. 9.14 Six carbamates eluted in fraction III (portion of Carbofuran and Methiocarb in fraction II); 5.0ng of each compound; gas chromatographic column, 5% OV-210. (Reprinted with permission from Thompson et al. [18]. Copyright (1977) Springer Verlag, New York.)
Carbamate insecticides and herbicides
579
Coburn et al. [315] reported a procedure for the extraction and analysis of N-methylcarbamates in natural water. Their procedure involved the extraction at pH3–4 by solvent partitioning with methylene chloride, salting out with sodium sulphate, hydrolysis of the extracts by methanolic potassium hydroxide to form the corresponding phenols, reextraction of the phenols at a pH of 2 or lower with methylene chloride, chemical derivatization with pentafluorobenzyl bromide to form the ether derivatives, clean-up on a silica gel microcolumn, and analysis by gas chromatography with electron-capture detection. Their recoveries were 87 to 98% for several carbamates, namely Propoxur, Carbofuran, 3ketocarbofuran, Metmercapturon, Carbaryl and Mobam. The determination of Mexacarbate and Aminocarb was not possible with this procedure as the phenolic products obtained on hydrolysis were not extracted from the acidified media. Cohen et al. [309] used an electron-capture gas chromatograph to determine carbamate insecticides as their 2,4-dinitrophenyl derivatives in water. After isolation from the sample material by the appropriate extraction method, the carbamates were reacted with 1-fluoro-2,4dinitrobenzene. The resulting 2,4-dinitrophenyl ethers were determined by subjecting their solution in hexane to gas chromatography on a glass column (140cm × 15mm) packed with 1% Epikote 1001 on AW-DMCS Chromosorb G (60–80 mesh) operated at 211°C; nitrogen was used as carrier gas. The recovery of carbamates added to water was in the range of 82–100% except for Carbaryl (63%) and Butacarb (4–65%). The limit of determination of carbamates in water was 0.005mg L-1. Sundaram et al. [317] have described a rapid and sensitive analytical technique to quantify carbamate insecticides at nanogram levels using sorption on an Amberlite XAD-2 resin column and desorption followed by nitrogen–phosphorus–gas–liquid chromatographic analysis. The carbamates were extracted from natural water by percolation through a column of Amberlite XAD-2 followed by elution with ethyl acetate. The carbamate residues were then analysed directly. Recoveries between 86 and 108% were obtained for the following insecticides: Aminocarb (4dimethylamino-M-tolyl)N-methylcarbamate, Carbaryl (1-naphthyl-Nmethylcarbamate), Mexacarbate (4-dimethylcarbamate) and Propoxur (oisopropoxyphenol-N-methylcarbamate). Only 41–58% recovery was obtained for methomyl(S-methyl-N-(methylcarbamoyl)-oxy)thiocetimidate. The thermally labile carbamate herbicide Aldicarb has been determined by gas chromatography–mass spectrometry [318] and is facilitated by the use of a short capillary column. The limits of detection for Aldicarb oxime and Aldicarb nitrile, respectively, are 0.3ng, 1.3ng and 0.05ng. Zhang et al. [319] developed a gas chromatographic method for the quantification of the pesticide Aldicarb and its oxidative metabolics in well water. Modifications for the determination of Carbofuran, Oxamyl
580
Insecticides and herbicides
and Methomyl are also described. An open tubular column and a nitrogen phosphorus detector are used to quantify levels of Aldicarb, Aldicarb sulphoxide and Aldicarb sulphone as low as 1µg L-1 in a single injection procedure. Experiments were carried out for the determination of each metabolite in solution in distilled and well water. Recoveries ranged from 96 to 105% with no difference noted between recoveries from the distilled and well waters. 9.6.1.3 Thin-layer chromatography N-Methylcarbamate and N,N’-dimethylcarbamates were determined in water samples by hydrolyses with sodium bicarbonate and the resulting amines reacted with 4-chloro-7-nitrobenzo-2,1,3-oxadiazole in isobutyl methyl ketone solution to produce fluorescent derivatives [320]. These derivatives were separated to thin-layer chromatography on silica gel G or alumina with tetrahydrofuran–chloroform (1:49) as solvent. The fluoresence is then measured in situ (excitation at 436nm, emission at 528 and 537nm for the derivatives of methylamine and dimethylamine, respectively). The method was applied to natural water samples containing parts per 109 levels of carbamate. The disadvantage of the method is its inability to differentiate between carbamates of any one class. Thin-layer chromatographic methods have been described [321] for determining N-arylcarbamate and urea herbicides in natural water (Barban, Chloroprophan, Diuron, Fenuron, Fenuron TCA, Linuron, Monouron, Monouron TCA, Neburan, Propham, Siduron and Swep) and for determining O-arylcarbamate pesticides in water (Aminocarb, Carbaryl, Methiocarb, Mexacarbate and Propoxur). To determine N-arylcarbamates a measured volume of water sample is extracted with methylene chloride and the concentrated extract is cleaned up with a Florasil column. Appropriate fractions from the column are concentrated and portions are separated by thin-layer chromatography. The herbicides are hydrolysed to primary amines, which in turn are chemically converted to diazonium salts. The layer is sprayed with 1-naphthol and the products appear as coloured spots. Quantitative measurement is achieved by visually comparing the response of sample extracts to the responses of standards on the same thinlayer plate. Direct interferences may be encountered from aromatic amines that may be present in the sample. To determine O-arylcarbamate insecticides a measured volume of water is extracted with methylene chloride. The concentrated extract is cleaned up with a Florasil column. Appropriate fractions from the column are concentrated and portions are separated by thin-layer chromatography. The carbamates are hydrolysed on the layer and the hydrolysis products are reacted with 2,6-dibromoquinone chlorimide to yield specific coloured products. Quantitative measurement is achieved by visually comparing the
Carbamate insecticides and herbicides
581
responses of sample extracts with the responses of standards on the same thin layer. Identification is confirmed by changing the pH of the layer and observing colour changes of the reaction products. Direct interferences may be encountered from phenols that may be present in the sample. These materials react with chromogenic reagent and yield reaction products similar to those of the carbamates. Carbofuran and its degradation products carbofuranphenol, 3-ketocarbofuran, 3-hydroxycarbofuranphenol, N-hydroxymethylcarbofuran and 3-hydroxycarbofuran, have been determined in water samples by thin-layer chromatography of ether extracts of the sample [322]. 9.6.1.4 High-performance liquid chromatography This technique has been discussed by various workers [323, 324]. Liquid chromatography with an electrochemical detector has been used [323] to estimate 2–7ppm of various carbamate insecticides in water. 9.6.1.5 Enzymic assay Yamani et al. [325] have described an automated method for pesticide detection based on immobilized butyrylcholinase derived from horse serum. Detection limits were in the range 1–3µg L-1. Rule et al. [326] have described a method for the determination of Carbofuran based on on-line immunoaffinity chromatography with coupled-column liquid chromatography–mass spectrometric detection. 9.6.1.6 Fluorimetric analysis Frei et al. [327] have described an in situ fluorimetric method for the determination of Carbaryl and 1-naphthol by thin-layer chromatography. The compounds are extracted from the water sample with dichloromethane, and are separated by thin-layer chromatography on activated silica gel (without binder) with chloroform as solvent (ascending); the R values are 0.43 and 0.60 for Carbaryl and its hydrolysis f product (1-naphthol), respectively. The chromatogram is sprayed with sodium hydroxide to convert Carbaryl into 1-naphthol and to make the spots fluorescent; they are located in UV radiation, marked and measured by fluorimetry on the plate at 488nm within 1.5h after spraying. Visual and instrumental detection limits are about 6 and 1ng per spot, respectively. Calibration graphs are rectilinear from 0.01 to at least 3µg per spot. Standard addition techniques can be used for small amounts (1– 5ng per spot). Recovery of Carbaryl added to water was 90–94%; that of 1-naphthol was only 50%. Other carbamates do not interfere with the determination.
582
Insecticides and herbicides
9.6.1.7 Miscellaneous Fernandez et al. [328] have described a flow-through sensor for the direct determination of down to microgram per litre levels of carbamate pesticide mixtures without chromatographic separation. Carbofuran, Propoxur and Carbaryl are particularly mentioned. Suzuki et al. [329] have discussed the determination of Benthiocarb (S-(4-chloroethyl)-N,N-diethylthiolcarbamate) herbicide in rivers and agricultural drainage. 9.6.1.8 Preconcentration XAD-7 cation-exchange resin has been used to preconcentrate Aminocarb prior to solvent extraction and estimation by gas chromatography using a nitrogen–phosphorus detector [330]. 9.6.2 Waste waters 9.6.2.1 Gas chromatography Onuska and Boos [331] have determined S-alkyl derivatives of N,N-dialkyl dithiocarbamate in waste water samples by a gas chromatographic–mass spectrometric method. Separation of the derivatives was carried out on a column of 10% Apiezon L on Varaport 80 operated at 250°C with helium as carrier gas and flame ionization detection. Separation was also achieved on a similar column as above, but with the temperature maintained at 190°C for 4–5min and then programmed at 4°C min-1 to 270°C, the helium flow rate being 28mL min-1. The effluent from the second column was examined by mass spectrometry–gas chromatography. 9.6.3 Trade effluents 9.6.3.1 High-performance liquid chromatography Hasler [332] has studied the rate of degradation of Zectron ((4-dimethylamine-3,5-xylyl)methylcarbamate) in alkaline water effluents. Highperformance liquid chromatography was used to identify xylenol as a degradation product of this herbicide. 9.6.4 Sewage effluents 9.6.4.1 High-performance liquid chromatography This procedure has been used to determine Aminocarb [333].
Substituted urea-type herbicides
583
Mullins and Kirkbright [334] determined sodium N-methyldithiocarbamate (Metham Sodium) and methylisothiocyanate fungicides in untreated sewage by high-performance liquid chromatography using a micellar mobile phase and UV detection. 9.7 Substituted urea-type herbicides 9.7.1 Natural waters Substituted urea herbicides with different substitutes are shown in Table 9.22.
Table 9.22 Substituted urea herbicides with different substituents
Key Buturon, 3-(4-chlorophenyl)-methyl-1 -(1-methylprop-2-ynyl)-urea Chlorobromuron, 3-(4-bromo-3-chlorophenyl)-1-methoxy-l-methylurea Chlorooxuron, 3-(4,4-chlorophenoxy)phenyl)-1,1 -dimethylurea Chlorotoluron, 3-(3-chlorotoluyl)-1,1-dimethylurea Diuron, 3-(3,4-dichlorophenyl)-1,1 -dimethylurea Fenuron, 1, 1-dimethyl-3-phenylurea Isoproturon, 3-(cumethyl)-1,1-dimethylurea Linuron, 3-(3,4-dichlorophenyl)-1-methoxy-l-methylurea Metabenzthiozuron, 3-(2-benzothiazolyl)-1,1-dimethylurea Metoxuron, 3-(3-chloro-6-methoxyphenyl)-1,1-dimethylurea Monolinuron, 3-(4-chlorophenyl)-1-methoxy-l-methylurea Monouron, 3-(4-chlorophenyl)-1,1-dimethylurea Neburon, 1-butyl-3-(dichlorophenyl)-1-methylurea Siduron, 1-(2-methylcyclohexyl-3)-phenylurea
584
Insecticides and herbicides
9.7.1.1 Gas chromatography Gas chromatography of phenylurea herbicides is difficult because of their ease of thermal decomposition. Procedures have been reported in which careful control of conditions allows these compounds to be gas chromatographed intact [335, 336]. Alternatively the phenylurea herbicides can be hydrolised to the corresponding substituted anilines which are then determined either by gas chromatography directly [337], or as derivatives [338], or colorimetrically after coupling with a suitable chromophore [342].
Gas chromatography using a nitrogen-specific detector has also been used [335, 339–341]. Gas chromatography of phenylurea herbicides is difficult because of their ease of thermal decomposition. Procedures have been reported in which careful control of conditions allows these compounds to be gas chromatographed intact [37, 335].
Substituted urea-type herbicides
585
In an electron-capture method for estimating Diuron (3-(3,4-dichlorophenyl)-1,1-dimethylurea) in surface waters, McKone and Hance [343, 344] extract the water sample (100mL) with dichloromethane (2 × 25mL) and the lower layers are combined and washed with water (5mL). After filtering through cotton wool, the organic solvent is evaporated under reduced pressure at 35°C. Saturated aqueous sodium chloride is added to the residue, the mixture is shaken, 2,2,4-trimethyl-pentane (5mL) is added, and the mixture is shaken again. An aliquot of the organic layer is then subjected to gas chromatography in a stainless steel column packed with 5% of E301 (methyl silicone) on Gas Chrom Q (60–80 mesh). Recoveries from controls and from pond, canal and river waters containing 0.001–1ppm of Diuron were about 94%, the coefficient of variation at the higher levels being about 70%. In a method described by Resales [345] 1-methoxy-1-methyl-3phenylurea herbicide is hydrolysed by phosphoric acid to give aniline and N-methodymethamine, which together with certain impurities in the commercial product are titrated with sodium nitrite solution. Aniline and certain byproducts are also determined separately by gas chromatography on a glass column (2m × 4mm) containing 10% of silicone oil OV-17 on Chromosorb Q (80–100 mesh) temperature programmed from 100 to 200°C at 4.5°C min-1, with thermal conductivity detection and helium carrier gas. Deleu et al. [346] has used two-dimensional thin-layer chromatography and gas chromatography to separate and identify 10 urea herbicides in concentrations down to 4µg L-1 in river waters (Fig. 9.15). Four different adsorbents were compared. The eluting solutions were: 1 2 3 4
diethyl ether-toluene (1:3) diethyl ether-toluene (2:1) equal volumes of 1 and 2 chloroform–nitromethane (3:1)
De Kok et al. [347] described two complete schemes for the analysis of 15 phenylurea herbicides and their degradation products, allowing the selective determination of each separate class of compounds and anilines in each other’s presence. They give descriptions of procedures involved in extraction; clean-up; catalytic hydrolysis of herbicides to anilines; derivatization with heptafluorobutyric anhydride; and liquid or gas chromatographic fractionation and/or separation. 9.7.1.2 Thin-layer chromatography Guthrie et al. [348] mentioned the use of thin-layer chromatography for the determination of Diuron (3-(3,4-dichlorophenyl)-1-1-dimethylurea) in river waters.
586
Insecticides and herbicides
Fig. 9.15 Two-dimensional chromatogram of: 1 = Metoxuron; 2 = Fenuron; 3 = Monouron; 4 = Isoproturon; 5 = Chlorotoluron; 6 = Diuron; 7 = Metabenzthiazuron; 8 = Neburon; 9 = Linuron, Chlorobromuron; 10 = Buturon A. (Reprinted with permission from Deleu et al. [346]. Copyright (1977) Elsevier Science Publishers BV.)
9.7.1.3 High-performance liquid chromatography Sulfonylureas form a class of herbicides introduced in the 1980s. From a chemical point of view sulphonylureas are labile and weakly acidic compounds. Compared to older herbicides, sulphonylurea herbicides have much lower use rates and are more rapidly degraded in soil. When present, very low concentrations (low parts per trillion region) of these herbicides in environmental waters are then to be expected. For this reason and because of the chemical characteristics cited above, monitoring of these herbicides in water is a particularly challenging problem. Although various separation techniques, such as gas chromatography–mass spectrometry of sulphonylurea derivatives [472], super-critical fluid chromatography [473] and capillary electrophoresis [474, 475], have been proposed to analyse sulphonylurea in various matrices, liquid chromatography is the technique of choice [476–481]. Liquid chromatographic methods have become even more attractive since the introduction of robust and sensitive devices, such as thermospray and electrospray, to interface liquid chromatography to mass spectrometry. Volmer et al. [482] evaluated the performances of both thermospray and electrospray ion sources for determining trace levels of sulphonylurea herbicides in
Substituted urea-type herbicides
587
water. In terms of specificity, they concluded that the latter source, in combination with a tandem mass spectrometer, was superior to the former one. Smith and Lord [349] have used liquid chromatography for the determination of Chlorotoluron (3-(3-chlorotoluyl)-1,1-dimethylurea) residues but report that Diuron and Monouron interfere in the chromatographic system used. Farrington et al. [350] have described high-performance liquid chromatography procedures for the determination of residues of the following phenylurea herbicides in water: • • • • • • • •
Chlorobromuron (3-(4-bromo-3-chlorophenyl)-1-methoxy-1-methylurea) Chlorooxuron (3-(4,4-chlorophenoxy)phenyl)-1,1-dimethylurea) Chlorotoluron (3-(3-chlorotoluyl)-1,1-dimethylurea) Diuron (3-(3,4-dichlorophenyl)-1,1-dimethylurea) Linuron (3-(3,4-dichlorophenyl)-1-methoxy-1-methylurea) Metobromuron Monolinuron (3-(4-chlorophenyl)-1-methoxy-1-methylurea) Monouron (3-(4-chlorophenyl)-1,1-dimethylurea)
Typical chromatograms, obtained from extracts of river waters, are shown in Fig. 9.16. The lower limits of detection were estimated to be 0.01ppm for river water; below these levels both coextractives and signal noise interfere. Care was taken during the evaporation of solutions of urons in methanol to ensure that the temperature did not rise above 55°C as at higher temperatures degradation occurs. A cleanup procedure was required to remove coextractives [351]. Lawrence and Turton [352] have listed high-performance liquid chromatographic data for 166 pesticides and related compounds. Included are chromatographic conditions such as type of packing material, column dimensions and mobile phase composition. Elution volumes or capacity factors are listed as measures of compound retention. The types of substrate and detectors used for analyses are included; where UV detectors were employed, wavelengths are given. Di Corcia et al. [483] have discussed methods for the trace analysis of sulphonylurea herbicides in potable waters. Seven commonly used sulphonylureas, viz. Thifensulphuron methyl, Metsulphuron methyl, Trisulphuron, Chlorosulphuron, Rimusulphuron, Tribenzuron methyl and Bensulphuron methyl were extracted from water by off-line solid-phase extraction with a Carbograph 4 cartridge. Sulphonylurea herbicides were then isolated from both humic acids and neutral contaminants by differential
588
Insecticides and herbicides
Fig. 9.16 Typical chromatograms obtained from 5 L injections of river water extracts; (a) unfortified and (b) fortified with uron herbicides at 0.1mg kg-1. 1, Monouron; 2, Monolinuron; 3, Metobromuron; 4, Chlorotoluron; 5, Diuron; 6, Linuron; 7, Chlorobromuron, 8, Chlorooxuron. (Reprinted with permission from Farrington et al. [350]. Copyright (1977) Royal Society of Chemistry.)
elution. Analyte fractionation and quantification were performed by liquid chromatography with UV detection. Recoveries of sulphonylurea herbicides extracted from 4L of potable water (10ng L-1 spike level), 2L of ground water (50ng L-1 spike level) and 0.2L of river water (250ng L-1 spike level) were not lower than 94%. Depending on the particular sulphonylurea herbicides, method detection limits were 0.6–2ng L-1 in drinking water, 2–9ng L-1 in ground water and 13–40ng L-1 in river water. A preservation study of sulphonylurea herbicides stored on the Carbograph 4 cartridge was conducted. Over 2 weeks of cartridge storage, no significant analyte loss was observed when the cartridge was kept frozen. Comparing this method with one using a C -extraction cartridge, the former appeared to be superior to the 18 latter in terms of sensitivity and, chiefly, of selectivity. This method involves confirmatory analysis by liquid chromatography– electrospray–mass spectrometry instrumentation equipped with a single-quadruple mass filter. Mass spectrometer data acquisition was performed by a time-scheduled three-ion selected ion-monitoring programme. The necessary structure-significant fragment ions were obtained by controlled decomposition of sulphonylurea herbicides adduct ions after suitably adjusting the electrical field in the desolvation chamber. Under three-ion selected ion monitoring
Triazine herbicides
589
condition, limits of detection (S/N = 3) calculated from the ion current profiles of those fragment or parent ions giving the lowest S/N values ranged from between 0.5 (Tribenzuron methyl) and 3ng (Metasulphuron methyl, Thiafensulphuron methyl) injected into the liquid chromatographic column. 9.7.1.4 Miscellaneous Some of the urea herbicides have been separated on cellulose columns following hydrolysis to amines [353]. Farrington et al. [350] favour the high-performance liquid chromatographic approach to the identification and determination of phenylurea herbicides because of the obvious limitations of alternative methods such as direct gas chromatography, gas chromatography of hydrolysis products or derivatives, and spectrophotometry. They modified earlier work on liquid chromatographic methods [354–361]. 9.7.1.5 Preconcentration Aluminosilicates (Partisil ODS) have been used for the preconcentration of substituted phenylurea herbicides in natural and waste waters [362]. 9.8 Triazine herbicides 9.8.1 Natural waters Some triazine herbicides are listed below: • • • • • • • • • • • • •
Atrazine (2-chloro-4-ethylamino-6-isopropylamino-1,3,5-triazine) Propazine Simazine (2-chloro-4,6-bis-ethylamino-1,3,5-triazine) Prometon Prometryne Atraton (2-ethylamino-4-isopropylamino-6-methoxy-1,3,5-triazine) Ametryne (2-ethylamino-4-isopropylamino-6-methylthio-1,3,5triazine) Terbutryne Terbutylazine(4-tert-butylamino-2-chloro-5-ethylamino-1,3,5-triazine) GS26571 (2-amino-4-tert-butylamino-5-methoxy-1,3,5-triazine) GS30033 (2-amino-4-chloro-5-ethylamino-1,3,5-triazine) Terbumeton Secbumeton
590
Insecticides and herbicides
9.8.1.1 Gas chromatography Not unexpectedly, this is the method of choice for the analysis of herbicides. McKone et al. [363] compared gas chromatographic methods for the determination of Atrazine (2-chloro-4-ethylamino-6-isopropylamino-1,3,5-triazine), Ametryne (2-ethylamino-4-isopropylamino-6methylthio-1,3,5-triazine) and Terbutryne in water. The herbicides were extracted from water with dichloromethane and the dried extracts were evaporated to dryness at a temperature below 35°C. By gas chromatography on a glass column (1m × 4mm) of 2% neopentyl glycol succinate on Chromosorb W (80–100 mesh) operated at 195°C and with a RbBr-tipped flame ionization detector the three herbicides could be separated and 0.001ppm of each detected. This method was found to be superior to spectrophotometric and polarographic methods. Purkayastha and Cochrane [364] compared electron-capture and electrolytic conductivity detectors in the gas chromatographic determination of Prometon, Atraton (2-ethylamino-4-isopropylamino-6methoxy-1,3,5-triazine), Propazine, Atrazine (2-chloro-4-ethylamino-6isopropylamino-1,3,5-triazine), Prometryne, Simazine (2-chloro-4,6-bisethylamino-1,3,5-triazine) and Ametryne (2-ethylamino-4-isopropylamino-6-methylthio-1,3,5-triazine) in inland water samples. They found that the electrolytic conductivity detector seemed to have a wider application than a 63Ni electron-capture detector; use of the latter detector necessitated a clean-up stage for all the samples studied. The conductivity detector could be used in analysis of water without sample clean-up. Good recoveries of Atrazine added to water were obtained by extraction with dichloromethane. Ramsteiner et al. [365] compared alkali flame ionization, microcoulometric, flame photometric and electrolytic conductivity detectors for the determination of triazine herbicides in water. Methanol extracts were cleaned up on an alumina column and 12 herbicides were determined by gas chromatography with use of conventional columns containing 3% Carbowax 20M on 80–100 mesh Chromosorb G. Hormann et al. [366] monitored various European rivers for levels of Atrazine, Aimazine, Terbumeton and dealkylated metabolites GS26571 (2-amino-4-tertbutylamino-5-methoxy-1,3,5-triazine) and GS30033 (2amino-4-chloro-5-ethylamino-1,3,5-triazine). The compounds were extracted into dichloromethane and quantitated by gas chromatography with nitrogen-specific detection. Selected results were verified by gas chromatography with mass fragmentographic detection. The limit of detection was usually 0.4µg L-1. A gas chromatographic method has been issued by the US Environmental Protection Agency [367] for determining at the microgram per litre level the following herbicides in water and waste
Triazine herbicides
591
water: Ametryne, Atraton, Atrazine, Prometon, Prometryne, Propazine, Secbumeton, Simazine and Tertbutylazine. The method describes an efficient sample extraction procedure and provides, through use of column chromatography, a method for the elimination of non-pesticide interferences and the preparation of pesticide mixtures. Identification is made by nitrogen-specific gas chromatographic separation, and measurement is accomplished by the use of an electrolytic conductivity detector or a nitrogen-specific detector. Steinheimer and Brooks [368] developed a multiresidue method for the simultaneous determination of seven triazine herbicides in surface and ground water at a nominal detection limit of micrograms per litre. The technique uses solvent extraction, gas chromatographic separation and nitrogen-selective detection devices. Solid-phase extraction techniques using chromatographic grade silicas with chemically modified surfaces were examined as an alternative to liquid–liquid partition and evaluated using three natural water samples. Solid-phase extraction was found to provide rapid and efficient concentration with quantifiable recovery. Jahda and Marha [369] investigated the isolation of s-triazines from water using continuous steam-distillation extraction prior to gas–liquid chromatography. Recoveries of seven triazine herbicides, Propazine, Terbutylazine, Atrazine, Prometryne, Terbutryne, Desmetryne and Simazine, from water at pH values of 5.7 and 9 are reported. Recovery rates were independent of pH but generally improved with increase in time of steam-distillation extraction from 1 to 3h. Low recovery rates were obtained for Simazine. Atrazine only gave good recovery rates after 3h steam-distillation extraction. Lee and Stokker [370] have developed a multiresidue procedure for the quantitative determination of 10 triazines in natural waters by a gas chromatographic method using a nitrogen–phosphorus detector. Ametryne, Atraton, Atrazine, Cyanazine, Prometon, Prometryne, Propazine, Simazine, Simetone and Simetryne were used. All of them could be successfully quantified on both the Ultrabond 20M and 3% OV1 columns. Extraction was by methylene chloride and clean-up on Florasil. Recoveries of triazines at 10, 1.0 and 0.1µg L-1 were between 87 and 108% except for Simetone and Simetryne which were only 80% at 0.1µg L-1. The method was validated with Ontario lake water at the three experimental concentrations and with two other natural waters at 1µg L1 . The detection limit was 0.025µg L-1. 9.8.1.2 Gas chromatography–mass spectrometry Isotope dilution gas chromatography–mass spectrometry was used [371] to determine 0.1–1µg L -1 of Atrazine, Lindane, Diazinon and
592
Insecticides and herbicides
pentachlorophenol in natural water. An accuracy of 86% and a precision of 8% were demonstrated. Zangwei et al. [372] determined Atrazine in water at sub-ppt levels using solid-phase extraction and gas chromatography–high resolution mass spectrometry. They used a C -bonded cartridge, followed by 18 column chromatography on Florasil to remove interfering substances. 9.8.1.3 Thin-layer chromatography Zawadzka et al. [373] and Abbott [374] used thin-layer chromatography to determine Simazine, Atrazine and Prometryne herbicides in water. After extraction of a 250–1000mL sample (water) or a 10–100mL sample with dichloromethane or ethyl ether at pH9, the organic extract was condensed and applied to a column of basic aluminium oxide (activity III) and the herbicides were eluted with ether containing 0.5% of water. The eluate was condensed and applied to a layer of silica gel G impregnated with fluorescein. The chromatograms were developed with chloroform–acetone (9:1). The plates were dried, and the spots were located by spraying with 0.5% Brilliant Green (CI Basic Green 1) in acetone and exposing to bromine vapour. The plates were evaluated planimetrically. For samples containing 5–100µg of herbicide per litre the recoveries were between 83 and 97%, Thin-layer chromatography has also been used [375] to determine ring-labelled [14C] Ametryne in water. The herbicide was applied to silica gel plates as a methanol solution. After development the spots were located under ultraviolet radiation, removed, and treated with liquid scintillation solution for counting. The limit of detection was 5ng. Fishbein [376] has reviewed thin-layer, paper chromatographic, and gas and column chromatographic procedures for the determination of triazine herbicides. Sherma [377] determined triazine and chlorphenoxyacetic herbicides in natural water samples by solid-phase extraction and quantitative thinlayer chromatography. Down to 10µg L-1 herbicide could be determined with recoveries of 70–100%. 9.8.1.4 Enzyme immunoassay Bushway et al. [378] used enzyme immunoassay to determine Atrazine residues in natural waters in amounts down to 1µg L-1. Thurman et al. [379] compared enzyme linked immunoassay with gas chromatography using a mass spectrometric detector for the determination in natural waters of down to 0.2–2µg L -1 triazine herbicides including Atrazine, Ametryne and Propazine.
Triazine herbicides
593
9.8.1.5 High-resolution fast atom bombardment–mass spectrometry This method [380] has been employed to determine microgram per litre levels of Atrazine in dichloromethane extracts of natural waters. Between 6 and 94µg L-1 Atrazine were found in well water samples. 9.8.1.6 Miscellaneous Aaronson et al. [381] have studied the occurrence of Atrazine in a farm pond following a fish kill. Baker’s yeast cells (Saccharomices cerevisae) have been successfully immobilized on to silica gel and used in on-line isolation and trace enrichment of Desisopropylatrazine, Desethylatrazine, Hydroxyatrazine, Simazine, Cyanazine, Atrazine, Carabaryl, Propanil, Linuron and Fenamiphos [484]. Since humic and fulvic acids were not extracted, no clean-up was necessary. The pesticides were spiked at 0.1–1µg L-1 in tap water, ground water and sea water and were preconcentrated using online solid-phase extraction into a yeast immobilized on silica gel precolumn followed by liquid chromatography with diode-array detection. All the variables that affect the enrichment step, such as amount of yeast immobilized, dimensions of the precolumn, sample pH and preconcentration flow rate, were optimized. The degree of selectivity was evaluated by comparing the chromatograms obtained after on-line sample preconcentration on the yeast precolumn with those obtained by on-line solid-phase extraction using a precolumn filled with C material. The relative standard deviation for the whole procedure in 18 the determination of the selected pesticides at the 0.3µg L -1 concentration level ranged from 1 to 9%, depending on the pesticide and the type of water. Detection limits within the range 0.01– 0.5µg L-1 were obtained by percolating only 25mL of water sample without any additional clean-up step. 9.8.2 Sea water 9.8.2.1 Gas chromatography Wu et al. [382] carried out measurements of the enrichment of Atrazine on the microsurface water of an estuary. These authors used a microsurface water sampling technique with a 16 mesh stainless steel screen collecting bulk sampled from the top 100–150µm of the surface [383]. The enrichment of Atrazine in the microsurface varied from none to 110 times with the highest enrichment between mid September and late October.
594
Insecticides and herbicides
Atrazine concentration in the actual microsurface was estimated to vary in the range 150–8850µg L-1. 9.8.3 Waste water 9.8.3.1 Gas chromatography The gas chromatographic method described in Section 9.8.1.1 [367] has been applied to the determination of nine triazine type herbicides in waste waters. 9.8.4 Sewage effluents 9.8.4.1 Gas chromatography–mass fragmentography The gas chromatography–mass fragmentography procedure has been used by Karlhuber et al. [384] to identify apparent Atrazine residues in sewage water. The samples were cleaned up by the procedure of Ramsteiner et al. [365]. An aliquot of sewage water was neutralized and extracted with dichloromethane. The extract was further cleanedup by passage through an alumina column. After concentration, Atrazine was injected into the gas chromatograph equipped with a Coulson electrolytic conductivity detector. Injection of a 12.5µg sewage water aliquot into a gas chromatograph equipped with the nitrogen-specific Coulson electrolytic conductivity detector showed a peak with the same retention time as Atrazine. From a standard injection an apparent Atrazine concentration of 5.5mg L-1 in the sewage sample was calculated. The value found seemed extremely high and the sample was reinjected on the gas chromatograph with mass fragmentographic detection. No peak showed up in the chromatogram at the retention time of Atrazine, indicating that there was less than 0.01µg L-1 of Atrazine in the sewage water. The peak in the nitrogen-specific chromatogram, therefore, was a nitrogen-containing interference with the same retention time as Atrazine. Using a 4L water sample containing 2mg Atrazine 82–98% recovery was obtained using this procedure. 9.9 Phenoxyacetic acid herbicides Some phenoxyacetic acid herbicides are listed below: • • •
2,4-D ester and sodium salts (2,4-dichlorophenoxyacetic acid) 2,4-DP (Dichlorprop; 2,4-dichlorophenoxypropionic acid) 2,4-DB (4-(2-dichlorophenoxyl)-butyric acid)
Phenoxyacetic acid herbicides
• • • • • • • • •
595
MCPA (4-chloro-2-methylphenoxyacetic acid) MCPB (4-(4-chloro-2-methylphenoxyl)-butyric acid) Silvex (2(2,4,5-trichlorophenoxyl)-propionic acid) MCPP (Mecoprop; mixture of Mecoprop and 2-(2-chloro-4-methylphenoxy)-propionic acid) 2,4,5-T (2,4,5-trichlorophenoxyacetic acid) Dicamba Trifluralin (2-methoxy-3,6-dichlorobenzoic acid) Methoxychlor Fenoprop
9.9.1 Natural waters 9.9.1.1 Gas chromatography Devine et al. [385] adjust the water sample (1L) to pH2 with hydrochloric acid and extract it with benzene (100, 50 and 50mL). The extract is dried over sodium sulphate, concentrated to 0.1mL and methylated by the addition of diazomethane in ethyl ether (1mL). After 10min, the volume is reduced to about 0.1mL, acetone is added and an aliquot is analysed by gas chromatography on one of three columns: (1) 5% SE-30 on 60–80 mesh Chromosorb W at 175°C, (2) 2% QF-1 on 90–100 mesh Anakron ABS at 175°C or (3) 20% Carbowax 20M on 60–80 mesh Chromosorb W at 220°C. In each instance nitrogen is the carrier gas and detection is by electron capture. The minimum detectable amount of pesticide in water was 2 parts per 109 for MCPA (4-chloro-2-methyl-phenoxyacetic acid) and 0.01– 0.05 parts per 109 for 2,4-D (2,4-dichloro-phenoxyacetic acid) and its esters, 2,4,5-T (2,4,5-trichlorophenoxyacetic acid), Dicamba, Trifluralin (2methoxy-3,6-dichlorobenzoic acid), and Fenoprop. Recoveries were 50– 60% for MCPA and Dicamba and 80-95% for the other compounds. Croll [33] has given details of the use of back-flushing with electroncapture gas chromatography for the determination of phenoxyacetic acid type herbicides in water. Attention is paid to equalization of column resistance under operating and back-flushing conditions; base-line drift is thereby minimized. The system has been successfully used (with a variety of stationary phases, temperature ranging from 25 to 225°C and nitrogen flow rates from 25 to 200mL min-1. Colas et al. [386] have described methods for the separation and determination of down to about 1ppm of phenoxyalkanoic herbicides. Those present as salts or esters are hydrolysed by heating the water sample (1L) under reflux with sodium hydroxide for about 1h and the free acids are then extracted at pH2 with chloroform or
596
Insecticides and herbicides
dichloromethane (recoveries usually about 70%). After evaporation of the solution to dryness, the residual free acids are dissolved in acetone and treated with diazomethane and the methyl esters are analysed on a temperature-programmed column (1.5m × 6mm) containing 5% silicone DOW710 on Chromosorb W AW (45–60 mesh). Helium is used as carrier gas. For some separations other stationary phases are used. Typical results are presented for MCPA, MCPP and 2,4-D. Possible interference from phenols, chlorinated biphenyls and surfactants is discussed. Larose and Chau [387] state that owing to the similar retention times of several common phenoxyacetic acid type herbicides, the alkyl esters are subject to incorrect identification if several herbicides are present. Also, the sensitivity obtainable by means of electron-capture detection of the alkyl esters by some herbicides, such as MCPA and MCPB is very poor and therefore the method is generally not suitable for the determination of these compounds in water. In addition, the methyl ester of MCPA has a very short retention time close to the solvent front and is prone to interference from sample coextractives, which usually appear in this region. In fact the MCPA methyl ester often cannot be detected even at higher levels because of overlapping with coextraction peaks when the same gas chromatographic parameters as for the determination of organochlorine pesticides are used. Hence other derivatives have been considered. Carnac [388] studied a modified technique of dynamic distribution in liquid–liquid systems used for concentration of traces of organic substances in water. An organic solvent is placed on granules of copolymer of styrene and divinylbenzene. The technique has been used for determination of phenoxyalkanecarboxylic acids by gas–liquid chromatography with a flame ionization detector using chloroform as the solvent; the limit of detection is 5–10µg L-1. Chau and Terry [389, 390] have discussed the disadvantages of the gas chromatography of methyl esters produced by reaction with diazomethane and have developed the reaction conditions for forming 2-chloroethyl and pentafluorobenzyl esters of phenoxyacetic acids. Agemian and Chau [391] have reported a method for determining low levels of 4-chloro-2-methylphenoxyacetic acid and 4-(4-chloro-2methylphenoxy)-butyric acid in waste waters by derivatization with pentafluorobenzyl bromide. The increased sensitivity of the pentafluorobenzyl esters of these two herbicides over the 2-chloroethyl and methyl esters as well as their longer retention times make pentafluorobenzyl bromide the preferred reagent. These workers [391] used a gas–liquid chromatograph equipped with a nickel detector, a 6ft × 1/4in i.d. (1.8m × 6mm) coiled glass column and an automatic sampler connected to a computing integrator for data
Phenoxyacetic acid herbicides
597
processing. The column used was 3.5% w/w OV-101 and 5.5% w/w OV-210 on 80–100 mesh Chromosorb W, acid washed and treated with dimethylchlorosilane. The operating conditions were as follows: injector temperature 220°C, column temperature 220°C, detector temperature 300°C, carrier gas argon–methane (9:1) at a flow rate of 60mL min-1. Agemian and Chau [391] found that the few organochlorine pesticides that are eluted in the same fraction as the pentafluorobenzyl derivatives of the phenoxyacetic acid herbicides do not interfere because they have distinct retention times. Organophosphorus pesticides do not interfere. Twenty-four of the most widely used phenols either are eluted with the PCBs and organochlorine pesticides or have distinct retention times from those of the pentafluorobenzyl esters of the two herbicides. The whole of the above procedure, at a level of 0.5µg L-1 MCPA in 1L of distilled water, gave an average recovery of 75–80% with a coefficient of variation between 9 and 15%. Agemian and Chau [392] using the above method compared the penta-fluorobenzyl bromide reagent for forming esters and the boron trichlor-ide-2-chloroethanol and dicyclohexylcarbondiimide-2chloroethanol reagents for forming 2-chloroethyl esters of phenoxyacetic acid type herbicides, coupled with a complete solvent extraction system to obtain multiresidue methods for determining these compounds in natural waters at sub-microgram per litre levels. The nine herbicides studied by these workers were Dicamba, MCPA, MCPB, 2,4-DB, Picloram (4-amino-3,5,6-trichloropicolinic acid), 2,4-D, 2,4,5-T, Silvex and 2,4-DP. Lopez-Avila et al. [393] used isotope dilution gas chromatography– mass spectrometry to determine Dicamba and 2,4-D in natural waters, at the low microgram per litre level. Stable labelled isotopes are spiked into the sample before extraction and the ratio of unlabelled isotope was used to quantitate the unlabelled compounds. Average recoveries exceeded 84% and the relative standard deviation was better than 19%. Triclophon (3,5,6-trichloro-2-pyridyloxyacetic acid) has been determined [394] in river waters in amounts down to 5ng L-1 by an acid– catalyst esterification reaction of the herbicide with boron trifluoride– trifluoroethanol. Triclophon was first extracted from acidified river waters with diethyl ether and the resulting concentrate esterified under nitrogen at 80°C for 1h. The trifluoroethyl ester product was cleaned up by silica gel column chromatography and determined by gas chromatography with electron-capture detection. Recoveries from actual river waters were 90–93% with coefficients of variation of less than 4%. With only slight modifications at the clean-up stage, the proposed method was useful for the simultaneous determination of MCPA, 2,4-D and 2.4.5-T.
598
Insecticides and herbicides
9.9.1.2 Thin-layer chromatography Bogacka and Taylor [395] determined 2,4-D and MCPA herbicides in water using thin-layer chromatography. In this method a 1L sample of filtered water is treated with 50g of sodium chloride and 5mL of hydrochloric acid and the herbicides are extracted into ethyl ether (200, 100 and 100mL). The extract is dried with anhydrous sodium sulphate, concentrated to a few millilitres and passed through a column (180mm × 15mm) of silica acid with 90% methanol–acetic acid (9:1) as stationary phase and the herbicides are eluted with 150mL of light petroleum saturated with the methanol–acetic acid mixture. The first 30mL of eluate is rejected. The remaining eluate is evaporated to dryness, and the residue is dissolved in ether and concentrated to about 0.1mL before thin-layer chromatography on silica gel G–Keiselgel G (2:3) (activated for 30min at 120°C) with light petroleum–acetic acid– liquid paraffin (10:1:2) as solvent. The developed plates are air dried, sprayed with 5% silver nitrate solution and dried, then sprayed with 2M potassium hydroxide– formaldehyde (1:1), dried at 130–135°C for 30min, sprayed with nitric acid, and observed in UV illumination. For the determination the spots are compared with standards. Evaluation by the method of standard addition gave recoveries of 95.1% and 88.8% respectively with standard deviations of 14.2% and 14.3% for 2,4-D and MCPA respectively. These workers [396] also examined thin-layer chromatography of 2,4DP (Dichlorprop) and MCPP (mixture of Mecoprop and 2-(2-chloro-4methylphenoxy)propionic acid). In this method the ethyl ether extract of the sample is purified on a column of silicic acid and the herbicides are separated by thin-layer chromatography on silica gel– Kieselgel (2:3) with light petroleum–acetic acid–kerosene (10:1:2) as solvent. The sensitivity is 3µg of either compound per litre, the average recoveries of Dichlorprop and MCPP are 85.7% and 87.4%, respectively, and the corresponding standard deviations were 13.9% and 15.5%. Meinard [397] described a new chromogenic reagent for the detection of phenoxyacetic acid herbicides on thin-layer plates. The separated phenoxyacetic acids are detected (as violet spots on white background) by spraying the plate with a solution of chromotropic acid ((4g) in water (40g) and sulphuric acid (56g)) then heating at 160°C. The limits of detection for 2,4-D, 2,4,5-T and MCPA range from 0.05 to 0.2µg by spraying with silver nitrate reagent followed by exposure to UV radiation. 9.9.1.3 Paper electrophoresis Purkayastha [398] examined the applicability of paper electrophoresis to nine ionizable chlorinated phenoxyacetic acid type herbicides including
Phenoxyacetic acid herbicides
599
2,4-D, 2,4,5-T, MCPA, Fenoprop, Dicamba, Trifluralin (2-methoxy-3,6dichlorobenzic acid), and Picloram (4-amino-3,5,6-trichloropicolinic acid). Solutions were applied to paper moistened with pyridine–acetic acid buffer solution, pH3.7, 4.4, or 6.5, and a voltage of 2–4kV was applied. After 30min the paper was air-dried, sprayed with ammoniacal silver nitrate solution, and exposed to UV radiation. Experimental variation that increased the mobility of the spots included the applied voltage from 2 to 4kV (potential gradients of 50–100V cm-1), and adding a foreign electrolyte (e.g. potassium nitrate) to the buffer. Addition of methanol to the buffer resulted in decreased mobility as well as a variation in the relative mobilities of the compounds. 9.9.1.4 High-performance liquid chromatography Arjmand et al. [399] determined Dicamba using solid-phase extraction and ion-pair high-performance liquid chromatography. The detection limit was 1.6µg L-1. 9.9.1.5 Miscellaneous Marshall [400] used two methods for the infrared analysis of Dicamb– MCPA and Dicamba–2,4-D formulations. The ‘indirect’ method involved precipitation of the herbicides with hydrochloric acid and extraction with chloroform. The chloroform extract was evaporated to dryness, the residue was dissolved in acetone and the herbicides were determined by measuring infrared extinctions at the relevant wave-lengths. The ‘direct’ method involved dissolving the sample in acetone and measuring infrared extinctions. Although both methods gave good precision, the ‘indirect’ method was the more accurate. Bogacka [401] used 4-aminophenazone as a reagent for the spectrophotometric determination of phenoxyacetic acid herbicides (2,4D, Dichlorprop, MCPA) in water. The herbicides are extracted from an acidified 1L sample of water with ethyl ether. The extract is evaporated and the residue is eluted for 1h with 10g of pyridine hydrochloride at 207–210°C for 2,4-D, or at 225–230°C for Dichlorprop or MCPA. The resulting phenol derivative is steam distilled into aqueous ammonia (1M) extracted with light petroleum (after acidification of the distillate) and re-extracted into 0.05M aqueous ammonia for coupling with 4aminophenazone in the presence of potassium ferricyanide. The extinction of this solution is measured at 515, 505 or 515nm, for 2,4-D, Dichlorprop or MCPA respectively. The respective sensitivities are 20, 20 and 80µg L-1 of water and the corresponding standard deviations of the recovery are 4.0%, 8.5% and 3.5%, The method is not suitable for the determination of mixed herbicides.
600
Insecticides and herbicides
Liquid–liquid extraction Suffet [402] has evaluated liquid–liquid extraction techniques for separating phenoxyacetic acid herbicides from river water. He used the pvalue (defined as the fraction of the total solute that distributes itself in the non-polar phase of an equivolume solvent pair) concept in the development of equations, based on liquid–liquid extraction theory, relating the number of extractions and the water-to solvent ratios for the maximum recovery of the herbicide. Calculations show that a pesticide with a p-value of 0.90 or greater in an aqueous system can be 95% extracted from the aqueous phase by up to five extractions with a total volume of solvent up to 500mL. He confirmed his equations by measurements with 2,4-D. By the application of this concept to the simultaneous quantitative extraction of phenoxyacetic acid herbicides from water, Suffet [403] showed that the best solvents for 2,4-D and 2,4,5T and their butyl and isopropyl esters are ethyl ether or ethyl acetate (2,4D and esters) and benzene (2,4,5-T and esters). Thus a 90% recovery of 2,4-D from 1L of an aqueous solution is obtained by a two-stage serial extraction with 200 and 50mL of ethyl acetate under p-value conditions. Turbid samples should be filtered before extraction. The most commonly used solvents for extracting phenoxyalkanoic acids have been ethyl ether [404–407] and chloroform, although benzene has also been used [400, 408]. Benzene is highly toxic, and has a high boiling point and relatively low dielectric constant compared with the above solvents. This solvent gave consistently low extraction efficiencies for 2,4-D and Dicamba. This supports Suffet’s [403] p-value for 2,4-D of 0.195 for benzene compared with 0.996 and 0.990 for ethyl acetate and ethyl ether, respectively. Chloroform is also unsuitable because it is more toxic and has a lower dielectric constant than the other three solvents and its vapours cause anomalous responses when it is used near a gas chromatograph with an electron-capture detector. Derivatization–gas chromatography Chemical derivatization of phenoxyalkanoic acid herbicides has been used as a means for forming less polar and more volatile compounds for the gas–liquid chromatographic analysis. Alkyl esters [409–416] have been used extensively for this purpose. Examples of earlier work utilizing diazomethane for the formation of methylesters of phenoxyalkanoic acid herbicides are discussed by Devine et al. [385] and Colas et al. [386]. The conditions for the preparation of nitration, bromination and silylation using N,O-bis-trimethylsilylacetamide and 2-chloroethylation derivatives, and 1-bromomethyl-2,3,4,5,6-pentafluorobenzene
Miscellaneous herbicides
601
derivatives of 2,4-D and other herbicidal acids prior to gas chromatographic analysis, have been studied [389, 417]. 9.9.2 Waste waters 9.9.2.1 Gas chromatography Hill et al. [418] developed a multiresidue gas chromatographic method for the analysis of chlorophenoxy herbicides in waste waters and waste water sludges. The method, however, can only determine the acid form of chlorophenoxy herbicides. It was applicable to six herbicides, including 4-chorophenoxyacetic acid, 4-chloro-2-methylphenoxyacetic acid, 2,4-D (2,4-dichlorophenoxyacetic acid) and 2,4,5-TP (2,4,5trichlorophen-oxypropionic acid). The limit of detection of the chlorophenoxy herbicide was 5.0µg L-1 in an 11mL sewage sample for all the herbicides, except 2,4-dichlorophenoxybutyric acid. 9.10 Miscellaneous herbicides 9.10.1 Natural waters
Dichlorbenil (2,6-dichlorobenzonitrile) The persistence of this herbicide in a farm pond has been studied [419]. Dicamba (2-methoxy-3, 6-dichlorobenzoic acid) This herbicide is discussed in methods for its codetermination with phenoxyacetic acid herbicides. Norris and Montgomery [420] have described a procedure for the determination of traces of Dicamba and 2,4D in streams after forest spraying. Dicamba and its metabolites (3,6dichlorosalicylic acid and 5-hydroxydicamba) were determined gas chromatographically. For analysis a 500mL aliquot of stream water was acidified to pH1 with hydrochloric acid and extracted with three 150mL portions of diethyl ether. Ether extracts were concentrated to 20mL and methylated with diazomethane in ether. The ether extracts were then concentrated to 1mL and injected into a gas chromatograph equipped with a microcoulometric detector. The 1.8m × 6.25mm glass column was packed with 60–80 mesh Gas-Chrom Q coated with 6% OV-1. The retention time of Dicamba was 2.6min. Methylation converts 3,6dichlorosalicylic acid metabolite to Dicamba and the 5-hydroxy-dicamba metabolite which has a retention time of 6.6min.
602
Insecticides and herbicides
Picloram (4-amino-3,5,6-trichloropicolinic acid) Abbott et al. [421] described a pyrolysis unit for the determination of Picloram and other herbicides in water. The determination is effected by electron-capture–gas chromatography following thermal decarboxylation of the herbicide. Hall et al. [422] reported further on this method. The decarboxylation products are analysed on a column (5mm i.d.) the first 6in (15cm) of which is packed with Vycor chips (2–4mm), the next 3.5ft (107cm) with 3% of SE-30 on Chromosorb W (60–80 mesh) and then 2ft (60cm) with 10% of DC200 on Gas Chrom Q (60–80 mesh). The pyrolysis tube, which is packed with Vycor chips, is maintained at 385°C. The column is operated at 165°C with nitrogen as carrier gas (110mL min -1). The method when applied to ethyl ether extracts of water and soil gives recoveries of 93 ± 4 and 90 ± 5% respectively. Dennis et al. [423] have reported on the accumulation and persistence of Picloram in surface waters and bottom deposits. Pyrazon (5-amino-4-chloro-2-phenyl-3-pyridizone) This pre- and post-emergent herbicide has been determined in water by spectrophotometric, thin-layer chromatographic methods [424–427] and by high-performance liquid chromatography [428]. The highperformance liquid chromatographic method is described below as it illustrates very well the applicability of this technique to trace organics and analysis in water. Pyrazon was isolated from water samples (500mL) by rotary evaporation to dryness in vacuo, extraction of the solid residue with methanol (2 × 25mL) and further evaporation of the methanol extract (to approx. 2mL). Final concentration (to 0.5mL) was achieved by removal under a stream of nitrogen. The equipment used consisted of two Model 6000A solvent delivery systems and a Model 660 gradient former (Waters Associates) and a Model CE212 variable wavelength UV monitor (Cecil Instruments) operated at 270nm. Syringe injections were made through a stop-flow septumless injection port. The column (15cm × 7mm i.d.) was packed in an upward manner with Spherisorb-ODS by a slurry procedure using acetone as slurry medium. A linear gradient was established from two solvent mixtures consisting of (a) 10% methanol in 1% acetic acid in water and (b) 80% methanol in 0.1% acetic acid in water. The initial concentration was 35% (b) in (a) and the final concentration was 100% (b) with the gradient terminated after 20min. The flow rate was maintained at 2.0mL min-1 throughout the analysis.
Miscellaneous herbicides
603
Fig. 9.17 Current–potential graphs for Glyphosate nitrosamine after anionexchange treatment and nitrosation of various amounts of Glyphosate added to 1L of tap water. A, 0 (reagent blank); B, 35; C, 70; D, 140; E, 210 g added. (Reprinted with permission from Brønstad et al. [255]. Copyright (1976) Royal Society of Chemistry.)
Crathorne and Watts [428] determined the recovery efficiencies of Pyrazon from water by analysing samples of river water spiked at levels of 10, 50, 100 and 200µg L-1. Glyphosate (N-phosphonomethylglycine) This herbicide is manufactured by Monsanto, and marketed under the name Round-up. Brøstad et al. [255] have described a polarographic method for determining Glyphosate residues in natural waters based on the polarography of N-nitroso derivative. The Glyphosate nitrosamine has a single well-defined, differential pulse wave with a peak potential of -0.78V. Figure 9.17 shows a typical polarogram for single samples of tap water carried through the entire procedure after fortification at various levels. A Glyphosate concentration of 70µg L-1 gives a distinct Glyphosate nitrosamine wave.
604
Insecticides and herbicides
4-(chloromethylsulphenyl)bromobenzene Shiraishi and Otsuki [429] identified and determined 4-(chloromethylsulphenyl)bromobenzene herbicide in lake water using a combination of gas chromatography and mass spectrometry. Paraquat (1,1'-dimethyl-4,4-bipyridium chloride and Diquat (1,1'-ethylene-2,2-bipyridylium bromide) Calderbank and Youens [430] and Pope and Benner [431] have described a spectrophotometric method for the detemination of Paraquat in water in amounts down to 0.1ppm. Paraquat, Trifluralin and Diphenamid have also been determined gas chromatographically in water [432, 433]. Soderquist and Crosby [432] added to the water sample (100mL), sulphuric acid (3mL) and platinum dioxide (25mg) and hydrogen was bubbled through for 1h, whereby Paraquat is converted into 1,1'dimethyl-4,4'-bipiperidyl. This is extracted with dichloromethane (3 × 50mL) in the presence of 11mL of 50% sodium hydroxide solution and the combined extract is treated with 0.01N hydrochloric acid (4mL) and evaporated in a rotary evaporator at 50–55°C. The aqueous residue is transferred with 1mL of 0.01N hydrochloric acid to a 15mL screw-cap tube and shaken with 50% sodium hydroxide solution (0.5mL) and carbon disulphide (1mL). Aliquots of the carbon disulphide phase (1– 10µl) are injected on to a glass column (66ft × 0.125in (20m × 3mm)) packed with 10% Triton X-100 and 1% potassium hydroxide on AWDCMS Chromosorb G (70–80 mesh), and operated at 150°C with nitrogen as the carrier gas (30–40mL min-1) and flame ionization detection. The calibration graph is rectilinear for up to 1ppm of Paraquat. The limit of detection is 0.1ppm but recovery is only 36–43%, although reproducible. To determine Paraquat in agricultural run-off water Payne [434] separated the sediment from the sample (2L) by adding calcium chloride to aid flocculation, leaving the mixture overnight in a refrigerator for the sediment to settle. A 1L aliquot of the filtrate is extracted with dichloromethane. The dichloromethane extracts are concentrated by evaporation and the Trifluralin and Diphenamid are determined by direct injection, without further purification on to a glass column 6ft × 0.25in o.d. (1.8m × 6mm)) packed with 10% DC 200 on Gas Chrom Q and operated at 220°C with helium as carrier gas (100mL min1 ) and a Coulson electrolytic-conductivity detector (N mode). Paraquat is determined in the filtrate by a modification of a conventional colorimetric method. Recoveries of the three substances were between 82 and 95% from water.
Miscellaneous herbicides
605
Cannard and Criddle [433] have described a rapid pyrolysis–gas chromatography method for the simultaneous determination of Paraquat and Diquat in pond and river waters in amounts down to 0.001ppm. These workers emphasize the precautions necessary to avoid errors due to adsorption of the herbicides on to glassware.
Although other reactions occur that give smaller fragments, it will be apparent that the pyrolysis of both compounds produces few products with relative molecular masses comparable to those of the free bases, a feature which renders the method particularly suitable for both quantitative and qualitative analysis. However, for best results the procedure must be strictly adhered to. The detection limits for the method applied to river waters are governed by two main factors: the size of sample that can conveniently be introduced into the pyrolysis tube and the ability of the column to resolve the bipyridyl peaks from those due to other pyrolysis products. The most complex pattern obtained shows that no interference with Paraquat will occur, and that only slight interference with Diquat is likely. However a small Diquat pyrolysis peak can interfere to a slight extent with the 4,4'-bipyridyl peak derived from Paraquat but the value for Paraquat may be simply corrected when appropriate, as the size of the interfering peak is proportional to the size of the 2,2'-bipyridyl peak derived from Diquat. Coha [435] used the ring oven technique to estimate traces of Paraquat and Diquat in water. Morfamquat and Diquat have been determined by reduction at the dropping mercury electrode [436]. Zen et al. [485] used square-wave polarography at a perfluorinated ionomer–clay-modified electrode (MCME) to determine down to 0.5ppb of Paraquat in natural waters. The clay that showed the best performance for the fabrication of the electrode is nontronite (Swa-1, ferruginous smectite). The elctrochemical behaviour of Paraquat showed that the cathodic peak at -0.70V versus Ag/AgCl permits adequate quantification of the analyte. Linear calibration curves are obtained over the 0–80ppb range, with a detection limit of 0.5ppb in pH8 phosphate buffer solution for 4min preconcentration time.
606
Insecticides and herbicides
Thiabendazole Capitan et al. [437] determined down to 0.1mg L-1 Thiabendazole residues in natural waters using solid-phase spectrofluorometry. Other fluorescent insecticides or herbicides did not interfere in this procedure. 9.10.2 Sea water Thiabendazole The solid-phase spectrofluorometric procedure [437] discussed in Section 9.10.1 has been applied to the determination of thiabendazole in sea water. 9.10.3 Potable waters Glyphosate The polarographic method [255] described in Section 9.10.1 has been applied to the determination of Glyphosate in potable waters. Paraquat and Diquat In a method [438] for the preconcentration of Paraquat and Diquat from potable water the sample is passed through an ion-exchange column, followed by desorption, reduction with sodium dithionite and measurement of the reduced forms at 390nm for Paraquat and 379nm for Diquat. Carpenter et al. [486] have described a method for the assay of two metabolites of the herbicides dimethyl tetrachloroterephthalate, monomethyl tetrachloroterephthalate and tetrachloroterephthalic acid via high-performance liquid chromatography with ion pairing. Samples are analysed via direct injection, without preparation, and analyte detection is accomplished with an ultraviolet photodiode array detector. The metabolites are extracted from positive samples with a petroleum ether–diethyl ether mixture, derivatized with N,O-bis(trimethylsilyl)trifluoroacetamide, and confirmed by gas chromatography–mass spectrometry. The high-performance liquid chromatographic analysis of spiked drinking water samples yielded a recovery range of 92–106% with a mean recovery of 101% for tetrachloroterephthalic acid and a recovery range of 92–101% with a mean recovery of 96% for monomethyl tetrachloroterephthalate. The minimum detection limits for these two metabolites were 2.4 and 2.7µg L-1, respectively. In addition the gas chromatography–mass spectrometry analysis of spiked reagent water yielded mean recoveries of 91% for
Growth regulators
607
monomethyl tetra-chloroterephthalate and 86% for tetrachloroterephthalic acid. Twenty drinking water samples were split and analysed by the high-performance liquid chromatographic and the gas chromatographic–mass spectrometric methods and by US EPA method 515.1. Comparable results were obtained. The high-performance liquid chromatographic method, which is amenable to automation, typically allows for the analysis of up to 40 samples overnight. 9.10.4 Waste waters Thiabendazole The solid-phase spectrofluorometric method [437] described in Section 9.10.1 for the determination of Thiabendazole has been applied to waste waters. 9.11 Growth regulators 9.11.1 Natural waters 9.11.1.1 Gas chromatography Dalapon (2,2-dichloropropionic acid) Earlier gas chromatographic methods for determining this growth regulator include those of Getzendaner [439] and Frank and Demint [440]. The method of Getzendaner [439] is applicable to plant tissues and body fluid and doubtless to water samples. The sample was extracted with ethyl ether and the residue was analysed by gas chromatography on a glass column (4ft (1.2m) × 2mm) of 4% LAC-2R plus 0.5% of phosphoric acid on Gas Chrom S (60–80 mesh) at 100°C with nitrogen as carrier gas (85mL min-1) and electron capture detection. Recoveries of about 90% were obtained for 10ppm of the herbicide. The Frank and Demint [440] method is directly applicable to water samples. After addition of solid sodium chloride (340g L-1) and aqueous hydrochloric acid (1:1) to bring the pH to 1, the sample was extracted with ethyl ether and the organic layer was then extracted with 0.1M sodium bicarbonate (saturated with sodium chloride and adjusted with sodium hydroxide to pH8). The aqueous solution adjusted to pH1 with hydrochloric acid was extracted with ether and after evaporation of the ether to a small volume, Dalapon was esterified at room temperature by addition of diazomethane (0.5% solution in ether) and then applied to a stainless steel column (5ft × 0.125in (1.5m × 3mm)) packed with
608
Insecticides and herbicides
Chromosorb P (60–80 mesh) pretreated with hexamethyldisilazane and then coated with 10% FFAP. The column was operated at 140°C, with nitrogen carrier gas (30mL min-1) and electron-capture detection. The recovery of Dalapon ranged from 91 to 100%; the limit of detection was 0.1ng. Herbicides of the phenoxyacetic acid type did not interfere; trichloroacetic acid could be determined simultaneously with Dalapon. In a more recently published method (Van der Poll and de Vos [441]) for the determination of Dalapon in natural water and plant tissues the herbicide is first esterified with 3-phenolpropanol-1 then determined by electron-capture–gas chromatography. As little as 0.001mg Dalapon per litre of water can be determined by this method. These workers used a gas chromatograph with two 63Ni electron-capture detectors. The detectors were operated in the pulse mode at 50V. Two columns, both glass, were used to determine the ester. Column A was packed with 3% OV-1 and 2.7% OV-210 on Gas Chrom Q (80–100 mesh), column B with 1.8% OV-1 and 2.7% OV-210 on Gas Chrom Q (80–100 mesh). The carrier gas was nitrogen, flow rate 70mL min-1 on both columns. The temperatures of the column oven, injectors and detectors were 160, 205 and 275°C respectively. Recoveries of between 94 and 103% were obtained. The response of the detector was linear up to nanogram amounts of the 3-phenylpropyl ester of Dalapon injected. As little as 0.001mg Dalapon per litre natural water can be determined by this method. 9.11.1.2 Capillary isotachophoresis Stransky [442] investigated the possibility of determining the growth regulator Chloromequat, the quaternary cationic herbicides Diquat and Paraquat, and the triazine herbicides Atrazine, Simazine, Atraton, Prometryne, Demetryne and Methoprotryne in water extracts by capillary isotachophoresis. The more basic triazines could be determined directly using enforced isotachophoresis but very weak triazine bases had to be derivatized by nucleophilic substitution of chlorine by electron-donor or quaternary ammonium groups. Triazines could be quantitatively determined from a threshold value of about 15µg L-1. 9.12 Mixtures of pesticides and herbicides 9.12.1 Natural waters 9.12.1.1 Gas chromatography Cohen and Wheals [443] used a gas chromatograph equipped with an electron-capture detector to determine 10 substituted urea and carbamate
Mixtures of pesticides and herbicides
609
herbicides in river water, soil and plant materials in amounts down to 0.001–0.05ppm. The methods are applicable to those urea and carbamate herbicides that can be hydrolysed to yield an aromatic amine. A solution of the herbicide is first spotted on to a silica gel G plate together with herbicide standards (5–10µg) and developed with chloroform or hexane– acetone (5:1). The plate containing the separated herbicide or the free amines is sprayed with 1-fluoro-1,4-dinitrobenzene (4% in acetone) and heated at 190°C for 40min to produce the 2,4-dinitrophenyl derivative of the herbicide amine moiety. Acetone extracts of the areas of interest are subjected to gas chromatography on a column of 1% of XE-60 and 0.1% of Epikote 1001 on Chromosorb G (AE-DCMS) (60–80 mesh) at 215°C Erney [73] had described a photochemistry technique using ultraviolet irradiation followed by gas chromatography to confirm the identity of organochlorine insecticides and some herbicides. Kongovi and Grochowski [444] discussed the problems arising during the analysis of pesticides and herbicides by gas chromatography and electron-capture detection. During a routine run of pesticide standards (Lindane, Endrin and Methoxychlor) five peaks were obtained, and this led to a study of the Endrin molecule contaminants, and more specifically, to the decomposition of the Endrin molecule in relation to temperature (220–235°C) and nitrogen flow rate. Conclusions were eventually reached that the retention periods of compounds generally provide good criteria for identification, but that this was not always the case, particularly with esters of low-molecular-weight acids. Also the retention period alone does not serve as an absolute identifying criterion; other confirmation, e.g. mass spectroscopy, is required. Lee et al. [445] developed a multiresidue method with a low detection limit for 10 commonly used acid herbicides in natural waters. These herbicides were Dicamba, MCPA, 2,4-DP, 2,3,6-TBA, 2,4-D, Silvex, 2,4,5T, MCPB, 2,4,5-DB and Picloram. The method used solvent extraction and the formation of pentafluorobenzyl esters. The derivatives were quantified by capillary column gas chromatography with electroncapture detection. The detection limit was 0.05µg L-1. Recoveries of herbicides from spiked Ontario lake water (0.5–1.0µg L-1) were 73–108% except for Picloram recovery which was 59% at 0.1µg L-1. Ishibashi and Suzuki [446] simultaneously determined Chlormethoxynil, Bifenox and Butachlor in river water by XAD-2 resin extraction and high-resolution electron-capture gas chromatography. 9.12.1.2 Gas chromatography-mass spectrometry Yamato et al. [447, 448] have described a combination of gas chromatography with field desorption mass spectrometric analysis applied to Benzthiocarb (S-(4-chlorobenzyl-N,N-diethylthiolcarbamate),
610
Insecticides and herbicides
Oxidiazon (2-t-butyl-4-(2,4-dichloro-5-isopropoxyphenyl)-2-1,3,4oxadiazolin-5-one) and CNP (2,4,6-trichlorophenyl-4'-nitrophenylether) herbicides in water, and other environmental samples. They demonstrated the usefulness of this technique for screening unknown compounds in the natural environment. 9.12.1.3 Thin-layer chromatography Abbott and Wagstaff [449] of the Laboratory of the Government Chemist UK have described a thin-layer chromatographic method for the detection of 12 acidic herbicides and 19 nitrogenous herbicides (carbamates, substituted ureas, and triazines). Smith and Fitzpatrick [450] have also described a thin-layer method for the detection in water and soil of herbicide residues, including Atrazine, Barban, Diuron, Linuron, Monouron, Simazine, Trifluralin, Bromoxynil, Dalapon, Dicamba, MCPB, Mecoprop, Dicloram, 2,4-D, 2,4DB, Dichlorprop, 2,4,5-T and 2,3,6-trichlorobenzoic acid. Neutral and basic herbicides were extracted from water made alkaline with sodium hydroxide or from soil, with chloroform; extracts of soil were cleaned up on a basic alumina containing 15% of water. Acidic herbicides were extracted with ethyl ether from water acidified with hydrochloric acid or from an aqueous extract of soil prepared by treatment with 10% aqueous potassium chloride that was 0.05M in sodium hydroxide and filtration into 4M hydrochloric acid. The concentrated chloroform solution of neutral and basic herbicides was applied to a precoated silica gel plate containing a fluorescent indicator and a chromatogram was developed two-dimensionally with hexane– acetone (10:3) followed after drying by chloroform– nitromethane (1:1). The spots were detected in UV radiation. Atrazine, Barban, Diuron, Linuron, Monouron, Simazine and Trifluralin were successfully separated and were located as purple spots on a green fluorescent background. The ether extracts were dried over sodium sulphate, concentrated, and applied to a similar plate, which was developed two-dimensionally with chloroform–anhydrous acetic acid (19:1) followed after by drying by benzene–hexane–anhydrous acetic acid (5:10:2). The spots were detected by spraying with bromocresol green. Bromoxynil and (as the acids) Dalapon, Dicamba, MCPA, MCPB, Mecoprop, Dicloram, 2,4-D, 2,4-DB, Dichlorprop, 2,4,5-T and 2,3,6-trichlorobenzoic acid were seen as yellow spots on a blue background. The limits of detection were 1ppm in soil and 0.1ppm in natural water. In a further thin-layer chromatographic method for determining carbamate and urea herbicides in water at the parts per 109 level Frei et al. [451] extracted a 500mL sample with dichloromethane (2 × 50mL) and
Mixtures of pesticides and herbicides
611
evaporated the combined extract to 1mL at room temperature in a rotary evaporator and then to dryness at 40°C. The residue was dissolved in acetone (1 or 2 drops) and 0.5mL of sodium hydroxide and heated to 80°C for 30–40min, cooled and shaken with 0.2mL of hexane. 10µL of the hexane layer is applied to a 0.25mm layer of silica gel GCaSO , 0.2% dansyl chloride in acetone is applied to the sample spot, 4 and the chromatogram developed by the ascending technique with benzene–triethylamine–acetone (75:24:1). The plate is sprayed with 20% triethanolamine in isopropyl alcohol or 20% liquid paraffin in toluene, then dried. The fluorescence of the spots of the dansyl derivatives of the aniline moieties is measured in situ. Results are reported for carbamate pesticides, e.g. Propham, Chloropropham and Barban, and the urea pesticides, Linuron, Diuron, Chlorbromuron and Fluometuron; detection limits are about 1ng. Two-dimensional chromatography was used to eliminate interference. 9.12.1.4 High-performance liquid chromatography Miles and Moye [452] resolved several classes of pesticides by highperformance liquid chromatography and detected them by fluorescence after post-column UV photolysis with or without prior conversion to ophthalaldehyde-2-mercaptoethanol derivatives. In the presence of ophthalaldehyde-2-mercaptoethanol fluorescence labelling reagent, most carbamates, carbamoyloximes, carbamothioic acids and substituted ureas gave sensitive responses, whereas dithiocarbamates, phenylamides and phenylcarbamates gave varied responses. In the absence of ophthalaldehyde-2-mercaptoethanol reagent, strong fluorescence was observed following photolysis of several substituted aromatic pesticides. Detection limits for Aldicarb sulphoxide, Aldicarb, Propoxur, Thiram and Neburon, representing several classes of pesticides were 2.5, 2.3, 3.3, 3.8 and 2.0g L -1 respectively. The relative fluorescence (compared to equimolar methylamine) of some 50 pesticides from 10 different classes, in three solvents, and in the presence of o-phthalaldehyde-2mercaptoethanol were recorded. Fluorescence responses were significantly affected by the choice of solvent. Marvin et al. [453] have described an automated high-performance liquid chromatographic system using solid phase extraction for the analysis of mixtures of pesticides in waters. Pesticides determined included Propoxur, Carbofuran, Propham, Captan, Chloropropham, Barban and Butyrate. A multiresidue method for determining pesticides has been described [454] which uses a graphitized carbon black cartridge to remove pesticides from the water sample followed by liquid chromatographic analysis. Down to 0.003 to 0.07µg L -1 of 35 pesticides could be
612
Insecticides and herbicides
determined. Pesticides studied included Oxamyl, Methomyl, Phoxan, 2,4,5-trichlorophenoxyacetic acid (2,4,5-T), 2,4-DB and MCPB. Di Corcia et al. [455] used graphitized carbon black cartridges to extract polar pesticides prior to high-performance liquid chromatography. The cartridge was back flushed with the eluent to extract the pesticides prior to chromatography of the extract. Down to 0.01µg L -1 of 27 polar insecticides were determined including Dichlorvos, Methoate, Oxamyl, Methomyl. 9.12.1.5 High-performance liquid chromatography–mass spectrometry Bellar and Budde [456] applied this technique to the determination of 52 pesticides. Cappiello et al. [457] determined acidic, basic and neutral pesticides employing a liquid chromatography–mass spectrometry particle beam interface. Cresenzi et al. [458] determined 20 acidic pesticides including 2,4dinitrophenoxynilacetic acid, Mecoprop and Bromoxynil in river water using a benchtop electrospray liquid chromatograph–mass spectrometer. A reversed-phase LCC column was employed. The 18 mobile phase was 0.1M K HPO –0.2M Bu NF. Levels of 2.5–200ng of 2 4 4 each pesticide could be determined by this technique with a recovery of >85%. 9.12.1.6 Mass spectrometry Schulten [459–461] have identified many pesticides and their metabolites using field desorption mass spectrometry. However, these workers did not apply this method to environmental samples. Lin Hung and Vyksner [462] determined Terbutryn, Aldicarb sulphone, Propoxur and Carbofuran in environmental waters using an electrospray interface combined with an ion-trap mass spectrometer. 9.12.1.7 Miscellaneous Johnson et al. [463] studied the solid-phase extraction of pesticides from water samples, e.g. Trifluralin, Simazine, Atrazine, Propazine, Diazinon, Parathion-methyl, Arochlor, Malathion, Parathion and Chlorpyrifos. 9.12.1.8 Preconcentration Dedek et al. [464] preconcentrated hydrophilic pesticides such as Methamidophos on an organic polymeric sorbent Wofatit Y77. The
References
613
pesticides were then desorbed with cold or hot methanol prior to analysis. References 1 Gorbech, S., Harring, R., Knauf, W. and Werner, H.J. Bulletin of Environmental Contamination and Toxicology 6 40 (1971) 2 Chau, A.S.Y. and Terry, K. J. Ass. Off. Anal. Chemists 55 1228 (1972) 3 Chau, A.S.Y. and Terry, K. Analytical Abstracts 24 2513 (1973) 4 Chau, A.S.Y. J. Ass. Off. Anal. Chemists 55 1232 (1972) 5 Chromopak News, No. 20 June (1979) 6 American Public Health Association. Standard Methods for the Examination of Water and Waste Water, 15th Edition, Method 509 A, P 293(1978) 7 Method S73. Supplement to the 15th Edition of Standard Methods for the Examination of Water and Waste Water. Selected Analytical Methods Approved and Cited by US Environmental Protection Agency, American Public Health Association, American Water Works Association, Water Pollution Control Federation (1978) 8 Shevchunk, I.A., Dubchenco, Y.G. and Nordenova, Y.S. Soviet J. Water Chemistry and Technology 7 73 (1985) 9 Kahanovitch, Y. and Lahav, N. Environmental Science and Technology 1974 8 762 ( ) 10 Suzuki, M., Yamato, Y. and Wanatabe, T. Environmental Science and Technology 11 1109 (1977) 11 Taylor, R., Bogacka, T. and Krasnicki, K. Chemia Analit. 19 73 (1974) 12 Weil, L. and Ernst, K.E. Gas-u. Wass. Fach. 112 184 (1971) 13 Simal, J., Crous Vidal, J., MariaCharro Arias, A., Boado, M.A., Diaz, R. and Vilas, D. An. Bromat. (Spain) 23 1 (1971) 14 Sackmauereva, M., Pal’usova, O. and Szokolay, A. Water Research 11 551 (1977) 15 Konrad, J.G., Pionke, H.B. and Chesters, G. Analyst (London) 94 490 (1969) 16 Pionke, H.B. Analytical Abstracts 17 2442 (1969) 17 Millar, J.D., Thomas, R.E. and Schattenberg, H.J. Analytical Chemistry 53 214 (1981) 18 Thompson, J.F., Reid, S.J. and Kantor, E.J. Arch. Environ. Contam. Toxicol. 6 143 (1977) 19 Johnson, L.C. Bulletin of Environmental Contamination and Toxicology 5 542 (1970) 20 Pionke, H.B., Konrad, J.G., Chesters, J.G. and Armstrong, D.E. Analyst (London) 93 363 (1968) 21 Kadoum, A.M. Bulletin of Environmental Contamination and Toxicology 3 65 (1968) 22 Lauren, G. Bulletin of Environmental Contamination and Toxicology 5 542 (1970) 23 Ballinger, D.G. Methods for Organic Pesticides in Water and Waste Water, US Govt. Printing Office, Washington D.C., 58 pp. (1971) 24 Brodtmann, N.V. Bulletin of Environmental Contamination and Toxicology 15 33 (1976) 25 Kawahara, F.K. Journal of the Water Pollution Control Federation 39 572 (1967) 26 Methods for Organic Pesticides in Water and Waste Water 1972-759-31/2113. Region 5–11U.S. Govt. Printing Office, Washington DC (1971)
614
Insecticides and herbicides
27 Stachel, B., Bactjer, K., Cetinskaya, M., Dueszepn, J., Lahl, U., Lierse, K., Thieman, W., Gobel, B. and Kozichi, R. Analytical Chemistry 53 1469 (1981) 28 Malaiyandi, M., Jenkins, E., Lee, P. and Bowron, M.J. Environmental Science and Health A20 219 (1985) 29 Method S73. Supplement to the 15th Edition of Standard Methods for the Examination of Water and Waste Water. Selected Analytical Methods Approved and Cited by the United States Environmental Protection Agency, American Public Health Association, American Water Works Association, Water Pollution Control Federation 30 Lamar, W.L., Goerlitz, D.F. and Law, L.M. Identification and Measurement of Chlorinated Organic Pesticides in Water by Electron-capture Gas Chromatography, US Geol. Survey Water-Supply Paper 1817-B, 12 pp. (1965) 31 Lamar, W.L., Goerlitz, D.F. and Law, L.M. Determination of Organic Pesticides in the Environment, Am. Chem. Soc. Advances in Chemistry, ser 60, p. 187(1966) 32 (US) Federal Water Pollution Control Administraton. FWPCA Method for Chlorinated Hydrocarbon Pesticides in Water and Wastewater, Cincinnati, Federal Water Pollution Control Adm. 29 pp. (1969) 33 Croll, B.T. Analyst (London) 96 810 (1971) 34 Law, L.M. and Goerlitz, D.F. J. Am. Oil Color Chem. Ass. 53 1276 (1969) 35 Brodtmann, N.V. J. Am. Wat. Wks Ass. 67 558 (1975) 36 Breidenbach, A.W. The Identification and Measurement of Chlorinated/ Hydrocarbon Pesticides in Surface Water, 1968-0-315-842, US Govt Printing Office, Washington DC 37 Kahn, L. and Wayman, C.H. Analytical Chemistry 36 1340 (1964) 38 Goldberg, M.C., Delong, L. and Sinclair, M. Analytical Chemistry 45 89 (1973) 39 Aspila, K.I., Carron, J.M. and Chau, A.S.Y. J. Ass. Off. Anal Chem. 60 1097 (1977) 40 Analytical Methods Manual, Inland Waters Directorate, Water Quality Branch, Ottawa, Ontario, Canada (1974) 41 Miles, J.R.W., Tu, C.M. and Harris, O.R. J. Econ. Entomol. 62 1334 (1969) 42 Chau, A.S.Y., Rosen, J.D. and Cochrane, W.P. Bull. Environ. Contam. Toxicol. 6 225 (1971) 43 Chau, A.S.Y. J. Am. Oil Color Chem. Ass. 57 585 (1974) 44 Thompson, J.F. (Ed.). Analysis of Pesticide Residues in Human and Environmental Samples, US Environmental Protection Agency, Research Triangle Park, NC, sec. 10A 12/2/74, p. 9(1974) 45 Eichelberger, J.W. and Lichtenberg, J.L. Environ Sci. Technol. 5 541 (1971) 46 Suzuki, M., Yamato, Y. and Wanatabe, T. Nippon Nogei Kagaku Kaishi 47 1 (1973) 47 Sackmauereva, M., Pal’usova, O. and Hluchan, E. Vod Hospod 10 267 (1972) 48 Szokolay, A., Uhnak, J. and Madaric, A.A. Chem. Zvesti 25 453 (1971) 49 Janak, J., Sackmauereva, M., Szokolay, A. and Madaric, A. Chem. Zsvesti 27 128 (1973) 50 Janak, J., Sackmauereva, M., Szokolay, A. and Pal’usova, O. Journal of Chromatography 91 545 (1974) 51 Sherma, J. and Shafik T.M. Arch. Environ. Contam. Toxicol. 3 55 (1975) 52 Kadoum, A.M. Bull. Environ. Contam. Toxicol. 3 247 (1968) 53 Askew, J. Ruzicka, J.H. and Wheals, B.B. Analyst (London) 94 275 (1969) 54 Herzel, F. Arch. Hyg. Bakteriol 154 18 (1970) 55 Ahling, B. and Jenson, S. Analytical Chemistry 42 1483 (1970)
References
615
56 Ripley, B.D., Wilkinson, R.J. and Chau, A.S.Y. J. Ass. Off. Anal. Chem. 57 1033 (1974) 57 ASTM Tentative Method of Test for Organochlorine Pesticides in Water Designation, D3086–72T (1972) 58 Harless, R.L., Harris, D.E., Sovocoal, W., Zehr, R.D., Wilson, N.K. and Oswald, E.O. Biochem. Mass Spectrometry 5 232 (1978) 59 Fehringer, N.V. and Westfall, J.E. Journal of Chromatography 57 397 (1971) 60 Chen, Jo-Yun T. and Dority, R.W. J. Am. Oil Color Chem. Ass. 55 15 (1972) 61 Hutzinger, O., Jamieson, W.D. and Safe, S. J. Am. Oil Color Chem. Ass. 54 178 (1971) 62 Kawatski, J.A. and Frasch, D.L. J. Ass. Off. Anal. Chem. 52 1108 (1960) 63 Arias, C, Vidal, A., Vidal, C. and Maria, J. An. Bromat. (Spain) 22 273 (1970) 64 Achari, R.G., Sandhu, S.S. and Warren, W.J. Bulletin of Environmental Contamination and Toxicology 13 94 (1975) 65 Bevenue, A., Kelley, T.W. and Hylin, J. Journal of Chromatography 54 71 (1971) 66 Suzuki, T., Nayayoshi, H. and Kashiwa, T. Agric. Biological Chemistry 38 279 (1974) 67 Suzuki, T. Analytical Abstracts 26 2374 (1974) 68 Armstrong, D.W. and Terrill, R.Q. Analytical Chemistry 51 2160 (1979) 69 Willmott, F.W. and Dolphin, R.J. Journal of Chromatographic Science 12 695 (1974) 70 Keith, L.H. and Alford, A.L. J. Ass. Off. Anal. Chemists 53 1018 (1970) 71 Sadar, M.H. and Guilbauelt, G.G. Journal of Agricultural and Food Chemistry 19 357 (1971) 72 Chau, A.S.Y. J. Ass. Off. Anal. Chemists 55 519 (1972) 73 Erney, D.R. Anal Lett. 12 501 (1979) 74 Novikova, K.E. Zhur. Vses. Khim. Obshch. 18 562 (1973) 75 Laubereau, P.G. Korrespondens Abwasser 31 499 (1984) 76 US Environmental Protection Agency, Report No EPA-600-4-74, 1974T 108 (1974) 77 Woodham, R.L. and Collier, C.W. J. Ass. Off. Anal. Chem. 54 1117 (1971) 78 Baird, R.B., Carmona, L.G. and Kuo, C.L. Bull. Environ. Contam. Toxicol. 9 108 (1973) 79 Croll, B.T. Chemy Ind. 40 1295 (1970) 80 Wood, G.W. Anal. Abstr. 23 4219 (1972) 81 Levi, I. and Nowicki, T.W. Bull. Environ. Contam. Toxicol. 9 20 (1973) 82 Bergemann, H. and Hellman, H. Deutsch Gewasserkundliche Mitteilungen 24 31 (1981) 83Le Roy, M.L. and Goerlitz, D.F. J. Ass. Off. Anal. Chemists 53 1276 (1970) 84 Weil, L. and Quentin, K.E. Z für Wasser und Abwasser 7 147 (1974) 85 Balayanis, P.G. Journal of Chromatography 90 198 (1974) 86 Godefroot, M., Stechale, M., Sandra, P. and Verzele, M. J. High Resolution Chromatography, Chromatography Communications 5 75 (1982) 87 Griffith, K.R. and Craun, J.C. J. Ass. Off. Anal. Chemists 57 168 (1974) 88 Burnham A.K. Analytical Chemistry 44 139 (1972) 89 Richards, J.F. and Fritz, J.S. Talanta 21 91 (1974) 90 Mangani, F., Cresentini, G. and Bruner, F. Analytical Chemistry 53 1627 (1981) 91 Bagnati, R., Bentenati, E., Davoli, E. and Fanelli, R. Chromosphere 17 59 (1988) 92 Sherma, J. Journal of Liquid Chromatography 11 2121 (1988) 93 Noy, T., Weiss, E., Herps, T., Von Crutchen, H. and Rijks, J. Journal of High Resolution Gas Chromatography, Chromatography Communications 11 181 (1988)
616
Insecticides and herbicides
94 Noroozian, E., Moris, F.A., Nielen, M.W.F., Frei, R.W., De Jong, G.J. and Brinkman, H.A.T. Journal of High Resolution Chromatography, Chromatography Communications 10 17 (1987) 95 Aue, W.A., Kapila, S. and Hastings, C.R. Journal of Chromatography 73 99 (1972) 96 A Guide to Marine Pollution, Chs 1,2 and 4 (E.D. Goldberg, editor). Gordon and Breach, New York (1972) 97 Braus, H., Middleton, F.M. and Watton, G. Analytical Chemistry 23 1160 (1951) 98 Middleton, F.M., Grant, W. and Rosen, A.A. Ind. Engng Chem. 48 268 (1956) 99 Ludzok, F.J., Middleton, F.M. and Ettinger, E.B. Sew. Ind. Wastes 30 662 (1958) 100 Polange, R.C. and Megergian, S. J. American Water Works Association 50 1214 (1958) 101 Middleton, EM. and Lichtenberg, J. J. Ind. Engng Chem. 52 99A (1960) 102 Standard Methods for the Examination of Water and Waste Water, 13th edn, APHA, AWWA, WPCF, Washington DC (1971) p. 103 103 Sproul, O.J. and Ryckman, D.W. WPCF 33 1188 (1961) 104 Rosen, A.A. and Middleton, F.M. Analytical Chemistry 31 1729 (1959) 105 Booth, R.L., English, J.N. and McDermott, G.N. J. Am. Wat. Wks Ass. 57 215 (1965) 106 Eichelberger, A. and Lichtenberg, J.J. J. Am. Wat. Wks Ass. 25 63 (1971) 107 Eichelberger, J.W. and Lichtenberg, J. J. Am. Wat. Wks Ass. 63 25 1971) 108 Brodtmann, N.V. J. American Water Works Association 67 558 (1975) 109 Weil, L. Gas-u.-Wassfach. 113 64 (1972) 110 Weil, L. Anal. Abst. 24 1259 (1973) 111 Beyermann, K. and Eckrich, W. Z. Anal. Chem. 265 1 (1974) 112 Shevchuk, L.A., Dubchenco, Y.G. and Naidenova, T.S. Soviet Journal of Water Chemistry Technology 7 73 (1985) 113 Taylor, R., Bogacka, T. and Krosnicki, K. Chemia Analit. 19 73 (1974) 114 Deubert, K.H. Bulletin of Environmental Contamination and Toxicology 5 379 (1970) 115 Devine, M.J. Agricultural and Food Chemistry 21 1095 (1973) 116 Fuka, T., Janda, V. and Triska, J. Vodni Hospodarstvi, Series B 33 245 (1983) 117 Lester, J.F. and Smiley, J.W. Bulletin of Environmental Contamination and Toxicology 7 43 (1972) 118 Poracro, P.J. and Shubiak, P. Analytical Chemistry 44 1865 (1972) 119 Chau, A.S.Y. and Terry, K. J. Ass. Off. Anal Chemists 57 394 (1974) 120 Woodham, D.W., Loftis, C.D. and Collie, C.J. Agricultural and Food Chemistry 20 163 (1972) 121 Fredeen, F.J.H., Saha, J.G. and Balba, M.H. Pesticides Monitoring Journal 8 241 (1975) 122 Hempel, D., Liebmann, R. and Hellwig, A. Forschr. Wasserchem. Ihrer. Grenzeb. 13 181 (1971) 123 Hughes, R.A. and Lee, G.F. Environmental Science and Technology 7 934 (1973) 124 Engst, R. and Knoll, R. Nahrung 17 837 (1973) 125 Wilson, A.J., Forester, J. and Knight, J. US Fish Wildlife Circular 355, 18– 20(1969), Centre for Estuaries and Research, Gulf Breeze, Florida US (1969) 126 Wilson, A.J. and Forester, J. J. Ass. Anal. Chemists 54 525 (1971) 127 Wilson, A.J. Bull. Environ. Contam. Toxicol. 15 515 (1976) 128 Picer, N., Picer, M. and Strohal P. Bulletin of Environmental Contamination and Toxicology 14 565 (1975)
References
617
129 Harvey, G.R. Absorption of chlorinated hydrocarbon from seawater by a crosslinked polymer. Woods Hole, Massachusetts, unpublished manuscript (1972) 130 Department of the Environment. Methods for the Examination of Waters and Associated Materials: Organochlorine Insecticides and Polychlorinated Biphenyls in Waters, 1978 Tentative Method (32313) HMSO, London, 28 pp. (1979) 131 Ronsen, E. American Laboratory 12 49 (1980) 132 Mosinska, K. Pr. Inst. Przem. Org. 1971 253 (1972) 133 Sherma, J. and Slobodien, R. Journal of Liquid Chromatography 7 2735 (1984) 134 Laseter, J.L., Dehion, I.R. and Remele, P.C. Analytical Chemistry 50 1169 (1978) 135 Kaiser, K.L.E. Science 185 523 (1974) 136 Andrade, P.S.L. and Wheeler, R.B. Bull Environ. Contam. Toxicol. 11 415 (1974) 137 Zitko, V. and Choi, P. PCB and Other Industrial Halogenated Hydrocarbons in the Environment. Fish. Res. Board Can. Technical Report No. 272, Biological Station, St. Andrews, N.B. (1971) 138 Oller, W.L. and Cramer, M.F. J. Chromat. Sci. B13 296 (1975) 139 Edwards, R. Chem. Ind (London) 1340 (1970) 140 Fishbein, L. Journal of Chromatography 58 345 (1972) 141 Schulte, E., Their, H.P. and Acker, L. Deut. Lebensm.-Rundsch. 72 229 (1976) 142 Goke, G. Deut. Lebensm.-Rundsch. 71 309 (1975) 143 Ahling, B. and Jensen, S. Analytical Chemistry 42 1483 (1970) 144 Dolan, J.W. and Hall, R.C. Analytical Chemistry 45 2198 (1973) 145 Musty, P.R. and Nickless, G. Journal of Chromatography 89 185 (1974) 146 Elder, G. Journal of Chromatography 121 269 (1976) 147 Södergren, A. Journal of Chromatography 160 271 (1978) 148 Södergren, A. Journal of Chromatography 71 532 (1972) 149 Blomberg, L. Chromatographia 8 324 (1975) 150 Bacaloni, A., Goretti, G., Lagano, A. and Petronio, B.M. Journal of Chromatography 175 169 (1979) 151 Thomas, O. Analytical Chemistry 62 1667 (1990) 152 Leoni, V. Journal of Chromatography 62 63 (1971) 153 Bavel, B., Jaaremo, M., Karisson, L. and Lindstream, G. Analytical Chemistry 68 1279 (1996) 154 Farwell, S.O., Beland, F.A. and Geer, R.D. Bulletin of Environmental Contamination and Toxicology 10 157 (1973) 155 Leoni, V., Pucetti, G., Columbo, R.J. and Ovidio, O. Journal of Chromatography 125 399 (1976) 156 Leoni, V., Pucetti, G. and Grella, A. Journal of Chromatography 106 119 (1975) 157 Gosink, T.A. Environmental Science and Technology 9 630 (1975) 158 Aue, W.A. Journal of Chromatography 13 329 (1975) 159 Bagnati, R., Bentenati, E., Davoli, E. and Fanelli, R. Chemosphere 17 59 (1988) 160 Noy, T, Weiss, E., Herps, T., Van Crutchen, H. and Rijks, J. J. High Resol. Chromatography, Chromatography Commun. 11 181 (1988) 161 Harvey, G.R. Report US Environmental Protection Agency, EPA-R2-73-177, 32 pp. (1973) 162 Niederschulte, U. and Ballschmiter, K.Z. Analytical Chemistry 269 360 (1974) 163 Junk, G.A., Richard, J.J., Greser, M.D., Witiaki, J.L., Anguello, M.D., Vick, R., Svec, H.J., Fritz, J.S. and Calder, G.V. Journal of Chromatography 99 745 (1974) 164 Kurtz. D.A. Bulletin of Environmental Contamination and Toxicology 17 391 (1977)
618
Insecticides and herbicides
165 Girenko, D.B., Klisenko, M.A. and Dishcholka, Y.K. Hydrobiological Journal 11 60 (1975) 166 Supplement to the 15th edition of Standard Methods for the Examination of Water and Waste Water. Selected Analytical Methods Approved and Cited by the US Environmental Protection Agency, American Public Health Association, American Water Works Association, Water Pollution Control Federation, Method S78 for PCBs in water and waste water, Sept . (1978) 167 Goerlitz, D.F. and Law, L.M. Method for Chlorinated Hydrocarbons in Water and Waste Water, US Environmental Protection Agency, Interim method, p. 7Sept. (1978) 168 Canagay, A.B. and Levine, P.L. US Environmental Protection Agency, Cincinnati, Ohio. Report EPA 600/2-79-166 (1974), 100 pp . ( P22 CAR) (1979) 169 McIntyre, A.E., Perry, R. and Lester, J.N. Environmental Pollution (Series B) 2 223 (1981) 170 Dunn, W.J., Stalling, D.L., Schwartz, T.F., Hogan, J.W., Petty, J.D., Johannson, E. and Wold, S. Analytical Chemistry 56 1308 (1984) 171 Ericksen, M.D., Stanley, J.S., Turman, J.K., Going, J.E., Redford, D.P. and Heggen, D.E. Environmental Science and Technology 22 71 (1988) 172 Mattson, D.E. and Nygren, S. Journal of Chromatography 124 265 (1976) 173 Jenson, S., Johnels, A.G., Olsson, M. and Otterlind, G. Ambio. Spec. Rep. 1 71 (1972) 174 Determination of Polychlorobiphenyls and Chlorinated Insecticides in Sewage Sludge, Association of Official Analytical Chemists, Washington DC, 10th edn, p. 393(1965) 175 Erne, K. Acta Pharmacol. Toxicol. 14 158 (1958) 176 Bartlett, J.K. and Skoog, D.A. Analytical Chemistry 26 1008 (1954) 177 Zimmerli, B., Sulser, H. and Marek, B. Mitt. Geb. Lebensmittelunters. Hyg. 62 60 (1971) 178 Cochrane, W.P. and Maybury, R.B. J. Ass. Off. Anal. Chem. 56 1324 (1973) 179 Miller, G.A. and Wells, C.E. J. Ass. Off. Anal. Chem. 52 548 (1969) 180 Jensen, S., Renberg, L. and Reutergard, L. Analytical Chemistry 49 316 (1977) 181 Pearson, J.R., Aldrich, F.D. and Stone, A.W. J. Agric. Food Chem. 15 938 (1967) 182 Ahnoff, M. and Josefsson, B. Bull. Environ. Contam Toxicol. 13 159 (1975) 183 Goerlitz, D.F. and Law, L.H. Bull. Environ. Contam. Toxicol. 6 9 (1971) 184 McIntyre, A.E., Perry, R. and Lester, J.N. Environ. Technol. Lett. 1 57 (1980) 185 McIntyre, A.E., Lester, J.N. and Perry, R. Analysis of Organic Substances of Concern in Sewage Sludge, final report to the Department of the Environment for contracts DGR/480/66 and DGR/480/240, pp. 42–45, Imperial College, London, UK Nov. (1979) 186 Holden, A.V. and Marsden, K. Journal of Chromatography 44 481 (1969) 187 Erikson, M.D. and Pellizzari, E.D. Bulletin of Environmental Contamination and Toxicology 22 688 (1979) 188 Snyder, D. and Reinert, R. Bulletin of Environmental Contamination and Toxicology 6 385 (1971) 189 Keith, L. Analysis of Organic Compounds in Two Kraft Mill Waste Waters, EPA600/4-75-005 (1975) 190 Keith, L. Environ. Sci. Technol. 10 555 (1976) 191 Keith, L. Identification and Analysis of Organic Pollutants in Water, ed. L.H. Keith, Ann Arbor Science, Ann Arbor, MI, Ch. 36 (1976)
References
619
192 Erikson, M.D., Zweidinger, R.A., Michael, L.C. and Pellizzari, E.D. Environmental Monitoring Near Industrial Sites: Polychlorinated Naphthalenes, EPA-560/6-77-019 (1977) 193 Erikson, M.D., Michael, L.C., Zweidinger, R.A. and Pellizzari, E.D. Environ. Sci. Technol. 12 927 (1978) 194 Dube, D.J., Veith, G.D. and Lee, G.F. Wat. Pollut. Contr. Fed. 46 966 (1974) 195 Lawrence, J. and Tosine, H. Environ. Sci. Tecnol. 10 381 (1976) 196 Shannon, E.E., Ludwig, F.J. and Valdemanis, I. Environmental Canada Research Report No. 49, Ottawa, 35 pp. (1976) 197 Bergh, A.K. and Peoples, R.S. Sci. Total Environ. 8 197 (1977) 198 Harper, D.B., Smith, R.V. and Gotto, D.M. Environ. Pollut. 12 223 (1977) 199 Lawrence, J. and Tosine, H. Bull. Environ. Contam. Toxicol. 17 49 (1977) 200 Liur, D., Chawla, V.K. and Chau, A.S.Y. Proc. 9th Ann. Conf. On Trace Substances in Environ. Health, Univ. Missouri, Columbia, Missouri, p. 189(1975) 201 Venkataraman, S. and Sathyamurthy, V. Journal of Indian Water Works Association 11 351 (1979) 202 Glass, R.L. Analytical Chemistry 53 921 (1981) 203 Frobe, Z., Drevenkar, V., Stengl, B., and Stefanac, Z. Anal. Chim. Acta 206 299 (1988) 204 Burguera, J.L. and Burguera, M. Anal. Chim. Acta 179 497 (1986) 205 De Loach, H.K. and Hemphill, D.D. J. Ass. Off. Anal. Chem. 52 333 (1969) 206 De Loach, H.K. and Hemphill, D.D. J. Ass. Off. Anal. Chem. 53 1129 (1970) 207 Brazhnikov, V.V., Poshememsky Sadodynskii, I.K. and Chernjakin J. Journal of Chromatography 175 221 (1979) 208 Laws, E.P. and Webley, D.J. Analyst (London) 86 249 (1961) 209 Bates, J.A.R. Analyst (London) 90 453 (1965) 210 Brody, S.S. and Chancy, J.E. J. Gas Chromat. 4 42 (1966) 211 Bache, C.A. and Lisk, D.J. Analytical Chemistry 37 1477 (1965) 212 Hartmann, C.H. Aerograph Research Notes 1–6 (1966) 213 Kanazawa, J. and Kawahara, T. Nippon Nogei Kagaku Kaishi 40 178 (1966) 214 Guiffrida, L., Ives, N.F. and Bostwick, D.C. J. Ass. Off. Anal. Chem. 49 8 (1966) 215 Kamen, A.J. Gas Chromat. 3 336 (1965) 216 Claborn, H.V., Mann, H.D. and Vehler, D.D. J. Ass. Off. Anal. Chem. 51 1243 (1968) 217 Burchfield, H.P., Rhoades, J.W. and Wheeler, R.J. J. Agric. Food Chem. 13 511 (1965) 218 Burchfield, H.P., Johnson, D.E., Rhoades, J.W. and Wheeler, R.J. J. Gas Chromat. 3 28 (1965) 219 Ives, N.F. and Fuiffrida, L. J. Ass. Off. Anal Chem. 50 1 (1967) 220 Dressler, M. and Janak, J. Colln. Czech. Chem. Commun. 33 3970 (1968) 221 Wooley, D.E. Analytical Chemistry 40 210 (1968) 222 Novak, A.V. and Malmstadt, H.V. Analytical Chemistry 40 1108 (1968) 223 Bowman, M.C. and Beroza, M. Analytical Chemistry 40 1448 (1968) 224 Bowman, M.C., Beroza, M. and Hill, K.R. J. Ass. Off. Anal. Chem. 54 346 (1971) 225 Kamen, A. J. Chromat. Sci. 7 541 (1969) 226 Skolnick M. J. Chromat Sci. 8 462 (1970) 227 Grice, H.W., Yates, M.I. and David, D.J. J. Chromat. Sci. 9 90 (1970) 228 Craven, D.A. Analytical Chemistry 42 1679 (1970) 229 Svojanousky, V. and Nebola, R. Chemickchisty 67 295 (1973)
620
Insecticides and herbicides
230 Greenhalgh, R. and Cochrane, W.P. Journal of Chromatography 70 37 (1972) 231 Suffet, I.H. and Faust, S.D. Journal of Agricultural and Food Chemistry 20 52 (1972) 232 Bargnoux, H., Pepin, D., Chahard, J.L., Vedrine, F., Petit, J. and Berger, J.A. Analysis 5 170 (1977) 233 Supplement to the 15th edition of Standard Methods for the Examination of Water and Waste Water. Selected Analytical Methods Approved and Cited by the US Environmental Protection Agency, American Public Health Association, Method S51 (1978) 234 Verweij, A., Regenhardt, C.E.A and Boter, H.L. Chemosphere 8 115 (1979) 235 Blanchet, P.F. Journal of Chromatography 179 123 (1979) 236 Daughton, C.G., Crosby, D.G., Carnos, R.L. and Hseih, D.P.H. Journal of Agricultural and Food Chemistry 24 236 (1976) 237 Daughton, C.G., Cook, A.M. and Alexander, M. Analytical Chemistry 51 1949 (1979) 238 Bourne, S. J. Environ. Sci. Hlth B 13 75 (1978) 239 Mallet, V.N., Berkane, K. and Caissie, G.E. Journal of Chromatography 139 386 (1977) 240 Puijker, L.M., Veenedaal, G., Janssen, H.M.J. and Griepink, B. Fresenius Z. Analyt. Chemie 306 1 (1981) 241 Cassita, A. and Mallett, V.N. Chromatographia 16 305 (1984) 242 Sherma, J. and Boymel, J.L. Journal of Chromatography 247 201 (1982) 243 Barren, A., DePablo, J. and Hernandez, S. Anal. Chim. Acta 212 123 (1988) 244 Simon, U.A. Liquid Chromatography-Gas Chromatography 5 899 (1987) 245 Otsuki, A. and Takaku, T. Analytical Chemistry 51 833 (1979) 246 Thibeau, R.J., Van Haverbeke, L. and Brown, C.W. Applied Spectroscopy 32 98 (1978) 247 Rodica, T. Revta Chem. 20 259 (1969) 248 Zycinski, D. Roczn. Panst. Zakl. Hig. 22 189 (1971) 249 Jovanovic, D.A. and Prosie, Z. Acta. Pharm. 22 91 (1972) 250 Leoni, V. and Pucceti, G. Farmaco. Ed. Prat. 26 383 (1971) 251 Schutzmann, O. and Barthel, R. Analytical Abstracts 18 3473 (1966) 252 Kovacs, O. Analytical Abstracts 13 2016 (1966) 253 Bidleman, T.F., Nowlan, B. and Frei, R.W. Anal. Chim. Acta 60 13 (1972) 254 Guven, K.C. and Aktugla, A. Eczacilik Bult. 14 44 (1972) 255 Brøstad, J.O., Hâkon, O. and Freidstad, J. Analyst (London) 101 820 (1976) 256 Gauson, R.M., Robinson, D. W and Goodman, A. Environmental Science and Technology 7 1137 (1973) 257 Karlhuber, B.A. and Eberle, D. Analytical Chemistry 47 1094 (1975) 258 Goodson, L.H., Jacobs, W.B. and Davis, A. Analyt. Biochem. 51 362 (1973) 259 Bauman, O. Anal. Abstr. 15 5550 (1968) 260 Report US Environ. Prot. Agency EPA-R 2-72 -010, Midwest Research Institute (1972) 261 Sharma, S., Rathore, H.S. and Ahmed, S.R. Water Research 17 471 (1983) 262 Goosen, J.T.H. and Kloosterboer, J.G. Analytical Chemistry 50 707 (1978) 263 Vogler, P. Liminologica 7 309 (1970) 264 Volpe, G.G. and Mallet, V.N. International Environmental Analytical Chemistry 8 291 (1980) 265 Hindin, E., Horteen, M.J., May, D.S., Skrinde, R.T. and Dunstan, G.H. J. Am. Water Wks Assoc. 54 88 (1962) 266 Skinde, R.T., Caskey, J.W. and Ellespie, C.K. J. Am Water Wks Assoc. 54 1407 (1962)
References
621
267 Teasley, J.T. and Cax, W.S. J. Am. Water Wks Assoc. 55 8 (1963) 268 Moye, H.A. Journal of Chromatographic Science 13 268 (1975) 269 Paschal, D.C., Bicknell, R. and Dresbach, D. Analytical Chemistry 49 1551 (1977) 270 Wolfe, N.L., Zepp, R.G., Gordon, G.A., Baughman, G.L. and Clive, D.M. Environ. Sci. Technol. 11 88 (1977) 271 Rice, J.R. and Dishberger, H.J. J. Agric. Food Chem. 16 867 (1968) 272 Deutsch, M.E., Westlake, W.E. and Gunther, F.A. J. Agric. food Chem. 18 178 (1970) 273 Dale, W.E. and Miles, J.W. J. Agric Food Chem. 17 60 (1969) 274 Miller, C.W. and Funes, A. Journal of Chromatography 59 161 (1971) 275 Shank, M.T. Bull. Environ. Contam. Toxicol. 3 309 (1968) 276 Howe, L.H. and Petty, C.F. Journal of Agricultural and Food Chemistry 17 401 (1969) 277 Coburn, J.A., Valdamanis, J.A. and Chau, A.S.Y. J. Ass. Off. Anal. Chem. 60 224 (1977) 278 Berkane, K., Caissie, G.E. and Mallett, V.N. Proc. Symp. on Fenitrothion, NRC (Canada) Assoc. Comm. Sci. Crit. Environ. Quality, Report 16073, NRCC, Ottawa, p. 95 (1977) 279 Zitko, V. and Cunningham, T.D. Bull. Environ. Contam. Toxicol. 14 19 (1975) 280 Mallet, V.N., Brun, G.L., MacDonald, R.N. and Berkane, K. Journal of Chromatography 160 81 (1978) 281 Berkane, K., Caissie, G.E. and Mallet, V.N. Journal of Chromatography 139 386 (1977) 282 Zakrevsky, J.G. and Mallet, V.N. Journal of Chromatography 132 315 (1977) 283 Ripley, B.D., Hall, J.A. and Chau, A.S.Y. Environ Lett. 7 97 (1974) 284 Grift, N. and Lockhart, W.L. J. Ass. Off. Anal. Chem. 57 1282 (1974) 285 Zakrevsky, J.G. and Mallet, V.N. Journal of Chromatography 132 315 (1977) 286 NRC Associate Committee on Scientific Criteria for Environmental Quality, Fenitrothion : The Effects of its Use on Environmental Quality and its Chemistry, NRCC No. 14104, p. 106 (1975) 287 Mallet, V.N. and Brun, G.L. Bulletin of Environmental Contamination and Toxicology 12 739 (1974) 288 Brun, G.L. and Mallet, V.N. Journal of Chromatography 80 117 (1973) 289 Weber, K. Water Research 10 237 (1976) 290 Cranmer, M.F. and Peoples, A. Analytical Biochemistry 55 255 (1973) 291 Cranmer, M.F. and Peoples, A. Analytical Abstracts 22 1779 (1972) 292 Supplement to the 15th edition of Standard Methods for the Examination of Water and Waste Water. Selected Analytical Methods Approved and Cited by the US Environmental Protection Agency, American Public Health Association, American Water Works Association, Water Pollution Control Federation, Method S51, September (1978) 293 Schulten, H.R. and Sun S.E. Int. J. Environ. Anal. Chem. 10 247 (1981) 294 Farran, A, Figuerola, E. and De Pablo, J. Int. J. Environ. Analytical Chemistry 33 245 (1988) 295 McIntyre, A.E., Perry, R. and Lester, J.N. Bulletin of Environmental Contamination and Toxicology 26 116 (1981) 296 Kawano, Y. and Bevenue, A. J. Chromatography 72 51 (1972) 297 Bilikova, A. Vov. Hospod. 21 16 (1971) 298 Procopio, J.R., Excribino, M.T.S. and Hernandez, L.H. Fresenius Z. für Analytische Chemie 331 27 (1988)
622
Insecticides and herbicides
299 Bosyakova, E.N., Bukharbaeva, A.S., Utebekova, N.R. and Shabanov, I.M. Trudy Inst. Braev. Patal, M-VOZ. Dravookir Kaz SSR 16 28 (1969). Ref ZhKhim 19GD (1969) 16 Abstract No. 169-210 300 Handa, S.K. and Dikshit, A.K. Analyst (London) 104 1185 (1979) 301 Handa, S.K. J. Ass. Off. Anal. Chemists 71 51 (1988) 302 Kumar, R. and Sharma, C.B. Analytical Letters (London) 20 777 (1987) 303 Condo, D.P. and Janauer, G.E. Analyst (London) 112 102 (1987) 304 Garcia Sanchez, F., Cruces, A. and Blanco, C. Analytical Chemistry 58 73 (1986) 305 Miskus, R., Gordon, H.T. and George, D.A. J. Agric. Food Chem. 7 613 (1959) 306 Venesch, E.E. and Riveros, M.H.C.K. J. Ass. Off. Anal. Chem. 54 128 (1971) 307 Rangaswamy, J.R. and Majumder, S.K. J. Ass. Off. Anal. Chem 57 592 (1974) 308 Crosby, D.G. and Bowers, O. Journal of Argiculture and Food Chemistry 16 839 (1968) 309 Cohen, I.C., Norcup, J., Kuzicka, J.H.A and Wheals, B.B. Journal of Chromatography 49 215 (1970) 310 Westlake, W.E., Monika, I. and Gunther, F.A. Bulletin of Environmental Contamination and Toxicology 8 109 (1972) 311 Reeves, R.G. and Woodham, D.W. Journal of Agriculture and Food Chemistry 22 76 (1974) 312 Rolls, J.W. and Cortes, A. J. Gas Chromatography 2 132 (1964) 313 Holden, E.R., Jones, W.M. and Beroze, M. J. Agric. Food Chem. 17 56 (1969) 314 Gutenmann, W.H. and Lisk, D.J. J. Agric. Food Chem. 3 48 (1965) 315 Coburn, J.A., Riplay, B.D and Chau, A.S.Y. J. Ass. Off. Anal. Chem. 59 188 (1976) 316 Lewis, D.L. and Paris, D.F. J. Agric. Food Chem. 22 148 (1974) 317 Sundaram, K.M.S., Szeto, S.Y. and Hindle, R. Journal of Chromatography 177 29 (1979) 318 Trehy, M.L., Yosf, R.A. and McCreary, J.J. Analytical Chemistry 56 1281 (1984) 319 Zhang, W.Z., Lemley, A.T. and Spalik, J. Journal of Chromatography 299 269 (1984) 320 Lawrence, J.F. and Frei, R.W. Analytical Chemistry 44 2046 (1972) 321 Supplement to the 15th Edition of Standard Methods for the Examination of Water and Waste Water. Selected Analytical Methods Approved and Cited by the US Environmental Protection Agency, American Public Health Association , American Waterworks Association , Water Pollution Control Federation , Sept. (1978). Methods S60 and S63. Methods for benzidine, chlorinated organic compounds, pentachlorophenol and pesticides in water and waste water (Interim, Pending issuance of methods for organic analysis of water and wastes, Sept. 1978), Environmental Protection Agency, Environmental Monitoring and Support Laboratory (EMSL) 322 Ya, C.C., Booth, G.H., Hanson, D.J. and Larsen, J.P. J. Agric. Food Chemistry 22 431 (1974) 323 Anderson, J.L. and Chesney, D.J. Analytical Chemistry 52 2156 (1980) 324 Anderson, J.L., Whiten, K.K., Brewster, J.D., Ou, T.Y. and Nonidez, W.K. Analytical Chemistry 57 1366 (1985) 325 Yamani, H., Iran-mihn, C. and Charanne, D. Sensors and Actuators 15 193 (1988) 326 Rule, G.S., Mordechei, A.V. and Henion, J. Analytical Chemistry 66 230 (1994) 327 Frei, R.W., Lawrence, J.F. and Belliveau, D.E. Z. Analyt. Chem. 254 271 (1971) 328 Fernandez, B., Linares, M.D., Luque de Castros, M.D. and Valearcel, M. Analytical Chemistry 63 1672 (1991)
References
623
329 Suzuki, M., Yamoto, Y. and Akiyama, T. Water Research 11 275 (1977) 330 Levesque, D. and Mallet, V.N. International Journal Environmental Analysis 16 139 (1983) 331 Onuska, F.I. and Boos, W.R. Analytical Chemistry 45 967 (1973) 332 Hasler, C.F. Bulletin of Environmental Contamination and Toxicology 12 599 (1974) 333 Brun, G.L. Bulletin of Environmental Contamination and Toxicology 24 886 (1980) 334 Mullins, F.G.P. and Kirkbright, G.F. Analyst (London) 112 701 (1987) 335 McKone, C.E. and Hance, R.J. J. Chromatography 36 234 (1968) 336 Khan, S.U., Greenhalgh, R. and Cochrane, W.P. Bull. Environ. Contam. Toxicol. 13 602 (1975) 337 Kirkland, J.J. Analytical Chemistry 34 428 (1962) 338 Lolke, H. Pest. Sci. 5 749 (1974) 339 Jarczyk, H.J. Pflanz. Nachr. Bayer 25 21 (1972) 340 Jarczyk, H. J. Pflanz. Nachr. Bayer 28 334 (1975) 341 Lowen, W.K., Bleidner, W.E., Kirkland, J.J., Pease, H.L. and Zweig, G. (Eds.) Analytical Methods for Pesticides, Plant Growth Regulators and Food Additives, Volume IV, Herbicides, Academic Press, New York, p. 157(1964) 342 Freistad, H. J. Ass. Off. Anal. Chem. 57 221 (1974) 343 McKone, C.E. and Hance, R.J. Bulletin of Environmental Contamination and Toxicology 4 31 (1960) 344 McKone, C.E. and Hance, R.J. Analytical Abstracts 17 3849 (1969) 345 Resales, J. Z. Analytical Chemistry 256 194 (1971) 346 Deleu, R., Barthelemy, J.P. and Copin, A. Journal of Chromatography 134 483 (1977) 347 De Kok, A., Vos, Y.J., Van Garderen, C, Se Jong, T., Van Opstal, M., Frei, R.W., Geerdink, R.B. and Brinkman, U.A.T. Journal of Chromatography 288 71 (1984) 348 Guthrie, R.K., Cherry, D.S. and Ferebee, R.W. Water Research Bulletin 10 304 (1974) 349 Smith, A.E. and Lord, K.A. Journal of Chromatography 107 407 (1975) 350 Farrington, D.S., Hopkins, R.G. and Ruzicka, J.H.A. Analyst (London) 102 377 (1977) 351 Onley, J.H. and Yip, G. Journal Ass. Off. Anal. Chemists 52 526 (1969) 352 Lawrence, J.F. and Turton, D. Journal of Chromatography 159 207 (1978) 353 Geissbuhler, H. and Gross, D. Journal of Chromatography 27 296 (1967) 354 Gotz, R. Facum Stadt Hygiene 29 10 (1978) 355 Shtivel, N.K., Gipikova, L.I. and Smirnova, Z.S. Soviet J. Water Chem. Tchnol. 7 66 (1985) 356 Mallevialle, J., Lefebvre, M., Roussau, C. and Sagbier, C. Rev. Fronc. Sci. L’eau 1 25 (1982) 357 Hunter, O. Environ. Sci. Technol. 9 241 (1975) 358 Stottmeister, E., Hendel, P. and Hermenau, H. Chemosphere 17 801 (1988) 359 Gomez-Belinchou, J.T. and Albaiges, J. Int. J. Environ. Anal. Chem. 30 183 (1987) 360 Schwarz, F.P. and Wasik, S.P. Analytical Chemistry 48 525 (1976) 361 Monarca, S., Causey, B.S. and Kirkbright, F.G. Water Research 13 503 (1979) 362 Senin, N.N., Filippov, V.S., Toliskina, N.F., Smolyaninov, G.A., Volkov, S.A. and Kukushkin, V.S. Journal of Chromatography 364 315 (1986) 363 McKone, C.E., Byast, T.M. and Hance, R.J. Analyst (London) 97 653 (1972)
624
Insecticides and herbicides
364 Purkayastha, R. and Cochrane, W.P. Journal of Agriculture and Food Chemistry 21 93 (1973) 365 Ramsteiner, K., Hoermann, W.D. and Eberle, D.O. J. Ass. Off. Anal. Chem. 57 192 (1974) 366 Hermann, W.D., Tournayre, J.C. and Egli, H. Pesticide Monitoring 13 128 (1979) 367 Method S69. Method for triazine pesticide in water and waste water. Methods for benzidine, chlorinated organic compounds, pentachlorophenol and pesticides in water and waste water (Interim, Pending Issuance of Methods for Organic Analysis of Water and Was 1978) 368 Steinheimer, T.R. and Brooks, M.G. International Journal of Environmental Analytical Chemists 17 97 (1984) 369 Jahda, V. and Marha, K. Journal of Chromatography 329 186 (1985) 370 Lee, H.B. and Stokker, Y.D. Journal of the Association of Official Analytical Chemists 69 568 (1986) 371 Lopez-Avila, V., Hirata, P., Kraska, S., Flanage, H. and Taylor, J.H. Analytical Chemistry 57 2797 (1985) 372 Zangwei, Lai, Sodagopa, O., Ramanujam, N.M., Giblin, D. and Gross, M.L. Analytical Chemistry 65 21 (1993) 373 Zawadzka, H., Adamezewska, M. and Elbanowska, H. Chemia Analit. 18 327 (1973) 374 Abbott, O. Anal. Abstr. 13 5917 (1966) 375 Frei, R.W. and Duffy, J.R. Mikrochim. Acta 3 480 (1969) 376 Fishbein, L. Chromat. Rev. 12 167 (1970) 377 Sherma, J. Journal of Chromatography 9 3433 (1986) 378 Bushway, R.J., Lekansi, S.T., Perkins, B., Savage, S.A. and Ferguson, B.S. Bulletin of Environmental Contamination and Toxicology 40 647 (1988) 379 Thurman, E.M., Meyer, M., Powes, M., Perry, C.A. and Schwab, A.B. Analytical Chemistry 62 2043 (1990) 380 Cadwell, K.A., Ramannjam, V.M., Cai, Z. and Gross, M.L. Analytical Chemistry 65 2372 (1993) 381 Aaronson, M.J., Kirby, K.W. and Tessari, J.D. Bulletin of Environmental Contamination and Toxicology 25 492 (1980) 382 Wu, T.L., Lambert, L., Hastings, D. and Banning, D. Bulletin Environ. Contam. Toxicol. 24 411 (1980) 383 Garrett, W.D. Limnol. Oceanography 10 602 (1965) 384 Karlhuber, B., Hermann, W. and Ramsteiner, K. Analytical Chemistry 47 2450 (1975) 385 Devine, J.N., Zweig, G. and Zip, B. Journal of Association of Official Analytical Chemists 52 187 (1969) 386 Colas, A., Lerenard, A. and Rover, J. J. Chim. Anal. 54 7 (1972) 387 Larose, R.H. and Chau, A.S.Y. Journal of Association of Official Analytical Chemists 56 1183 (1973) 388 Carnac, V.D. Zhurnal Analiticheskoi Khimmi 30 2444 (1975) 389 Chau, A.S.Y. and Terry, K. Journal of Association of Official Analytical Chemists 58 1294 (1975) 390 Chau, A.S.Y. and Terry, K. Journal of Association of Official Analytical Chemists 59 633 (1976) 391 Agemian, H. and Chau, A.S.Y. Journal of Association of Official Analytical Chemists 59 732 (1976) 392 Agemian, H. and Chau, A.S.Y. Journal of Association of Official Analytical Chemists 60 1070 (1977)
References
625
393 Lopez-Avila, V., Hirta, V., Kraska, S. and Taylor, J.H. Journal of Agriculture and Food Chemistry 34 530 (1986) 394 Tsukoika, T., Takeshita, R. and Murakami, T. Analyst (London) 111 145 (1986) 395 Bogacka, T. and Taylor, R. Chemia Analit. 15 143 (1970) 396 Bogacka, T. and Taylor, R. Chemia Analit. 16 215 (1971) 397 Meinard, C. Journal of Chromatography 61 173 (1971) 398 Purkayastha, R. Bulletin of Environmental Contamination and Toxicology 4 246 (1969) 399 Arjmand, M., Spittler, N.N. and Mumma, R.O. Journal of Agriculture and Food Chemistry 36 492 (1988) 400 Marshall, M. J. Association of Official Analytical Chemists 54 706 (1971) 401 Bogacka, T. Chemia Analit. 10 59 (1971) 402 Suffet, I.H. J. Agric. Food Chem. 21 288 (1973) 403 Suffet, I.H. J. Agric. Food Chem. 21 591 (1973) 404 Woodham, D.W., Mitchell, W.G., Loftis, C.D. and Collier, C.W. J. Agric. Food Chem. 19 186 (1971) 405 Bache, C.A., Lisk, D.J. and Loos, MA. J. Am. Oil Color Chem. Ass. 47 348 (1964) 406 Mattson, P.E. and Kirsten, W.J. J. Agric. Food Chem. 16 908 (1968) 407 Smith, A.E. and Fitzpatrick, A. Journal of Chromatography 57 303 (1971) 408 Devine, J.M., Zweig, G. and Zip, B. Journal of American Oil Color Chemists Association 52 187 (1969) 409 Meagher, W.R. J. Agric. Food Chem. 14 374 (1966) 410 McCone, C.E. and Hance, R.J. Journal of Chromatography 69 204 (1972) 411 Yip, G. J. Am. Oil Color Chemists Ass. 45 367 (1962) 412 Yip, G. J. Am. Oil Color Chemists Ass. 47 1116 (1964) 413 Suzuki, S.H. and Malina, M. J. Am. Oil Color Chemists Ass. 48 1164 (1965) 414 Goerlitz, D.F. and Lamar, W.L. Determination of Phenoxy Acid Herbicides in Water by Electron-Capture and Mircrocoulometric Gas Chromatography, Geological Survey Water-Supply Paper 1817-C, US Government Printing Office, Washington, DC (1967) 415 Yip, G. J. Am. Oil Color Chemists Ass. 54 966 (1971) 416 Howard, S.F. and Yip, G. J. Am. Oil Color Chemists Ass. 54 970 (1971) 417 Johnson, L.G. Journal of the Association of Official Analytical Chemists 56 1503 (1973) 418 Hill, N.P., McIntyre, A.E., Perry, R. and Lester, J.N. Int. Journal of Environmental Analytical Chemistry 15 107 (1983) 419 Rice, C.P., Sikka, H.C. and Lynch, R.S. Journal of Agricultural and Food Chemistry 22 533 (1974) 420 Norris, L.A. and Montgomery M.L. Bulletin of Environmental Contamination and Toxicology 13 1 (1975) 421 Abbott, S.D., Hall, R.C. and Giam, G.S. Journal of Chromatography 45 317 (1969) 422 Hall, R.C., Giam, G.S. and Merkle, M.G. Analytical Chemistry 42 423 (1970) 423 Dennis, D.S., Gillespie, W.H., Moxey, R.A. and Shaw, R. Arch. Environ. Contam. Toxicol 6 421 (1977) 424 Drescher, N. Methodensamminiung zur Ruckstandardsanalytik von Pflanzenschutsmitteln, Verlag Chimie, Weinheim (1972) 425 Drescher, N. Bestimmung der Ruckstande van Pyramin in Pflanze und Boden, 8– 9 January 1964 , BASF, Ludwigshafen am Rhein, p. 78(1964) 426 Zaborowska, W., Witkowska, I. and Kozak, H. Rocz. Panstu. Zakl. Hig. 24 735 (1973)
626
Insecticides and herbicides
427 Palusova, O., Sackmauerova, M. and Madarix, A. Journal of Chromatography 106 405 (1975) 428 Crathorne, B. and Watts, C.D. Journal of Chromatography 169 436 (1979) 429 Shiraishi, H. and Otsuki, A. Water Research 21 843 (1987) 430 Calderbank, A. and Youens, O. Analyst (London) 90 99 (1965) 431 Pope, J.D. and Benner, J.E. J. Ass. Off. Anal. Chem. 57 202 (1974) 432 Soderquist, C.J. and Crosby, D.G. Bull. Environ. Contam. Toxicol 8 363 (1972) 433 Cannard, A.J. and Griddle, W.J. Analyst (London) 100 848 (1975) 434 Payne, S.L. Journal of Agriculture and Food Chemistry 22 79 (1974) 435 Coha, O. Analytical Letters 2 623 (1969) 436 Volke, J. and Velkova, V. Collection Czech Chemical Communications 34 2037 (1969) 437 Capitan, F., Alonso, E., Avidad, R., Capitan-Valley, L.F. and Vilchez, J.L. Analytical Chemistry 65 1336 (1993) 438 Water Research Council. Methods for the Examination of Waters and Associated Materials, 1988 . Determination of Diquat and Paraquat in river and drinking waters, spectrophotometric methods, tentative, H.M. Stationery Office, London. (1987) 439 Getzenander, M.E. J. Ass. Off. Analytical Chemists 52 824 (1969) 440 Frank, P.A. and Demint, R.J. Environ. Sci. Technol. 3 69 (1969) 441 Van der Poll, J.M. and de Vos, R.H. Journal of Chromatography 187 244 (1980) 442 Stransky, Z. Journal of Chromatography 320 219 (1985) 443 Cohen, I.C. and Wheals, B.B. Journal of Chromatography 43 233 (1969) 444 Kongovi, R.R. and Grochowski, R. American Laboratory, p. 30(1981) 445 Lee, H.B., Stokker, D. and Chau, A.S.Y. Journal of Association of Official Analytical Chemists 69 557 (1986) 446 Ishibashi, M. and Suzuki, M. Journal of Chromatography 456 382 (1988) 447 Yamato, Y., Suzuki, M. and Wanatabe, T. Biochemical Mass Spectrometry 6 205 (1979) 448 Yamato, Y, Suzuki, M. and Wanatabe, T. J. Ass. Off. Anal. Chem. 61 1135 (1978) 449 Abbott, D.C. and Wagstaff, P.J. Journal of Chromatography 43 361 (1969) 450 Smith, A.E. and Fitzpatrick, A. Journal of Chromatography 57 303 (1971) 451 Frei, R.W., Lawrence, J.F. and Le Gay, D.S. Analyst (London) 98 9 (1973) 452 Miles, C.J. and Moye, H.A. Analytical Chemistry 60 220 (1988) 453 Marvin, C.H., Brindle, I.D., Hall, C.D. and Chiba, M. Analytical Chemistry 62 1495 (1990) 454 Jones, E.O. Analytical Chemistry 63 580 (1991) 455 Di Corcia, A., Samperi, R., Mariomini, A. and Stilluto, S. Analytical Chemistry 65 907 (1993) 456 Bellar, T.A. and Budde, W.L. Analytical Chemistry 60 2076 (1988) 457 Cappiello, A., Famiglini, G. and Bruner, F. Analytical Chemistry 66 1416 (1994) 458 Cresenzi C, Di Corcia, A., Marchese, S. and Sampori R. Analytical Chemistry 68 1968 (1996) 459 Schulten, H.R. and Beckey, D.H. J. Agric. Food Chem. 21 272 (1973) 460 Schulten, H.R., Prince, H., Beckey, H.D., Tomberg, W. and Korte, F. Chemosphere 2 22 (1979) 461 Schulten, H.R. J. Agric. Food. Chem. 24 743 (1976) 462 Lin Hung, Y. and Vyksner, R.D. Analytical Chemistry 65 451 (1993) 463 Johnson, W.E., Fendinger, N.J. and Plummer, J.R. Analytical Chemistry 63 1510 (1991)
References
627
464 Dedek, F., Wenzel, K.D., Luft, F., Obeelander, H. and Mothes, B. Fresenius Zeitschrift für Analytische Chemie 328 484 (1987) 465 Lacorte, S., Molina, C. and Barcelo, D. Anal. Chim. Acta 281 71 (1993) 466 Barcelo, D. Journal of Chromatography 643 117 (1993) 467 Barcelo, D., Mans, F.A., Geerdink, R.H., Frei, R.W. and Brinkman, U.A. Journal of Chromatography 394 65 (1987) 468 Locorte, S. and Barcelo, D. Analytical Chemistry 68 2464 (1996) 469 Russell Everett, W. and Rechnitz, G.A. Analytical Chemistry 70 807 (1998) 470 Molina, C., Honing, M. and Borelo, D. Analytical Chemistry 66 444 (1994) 471 Di Corcia, A., Crescenzi C. and Samperi, R. American Chemical Society, Journal of Environmental Chemistry. Preprints of papers presented at National Meeting Anaheim CA April 2–7th (1995) 472 Klaffenbach, P. and Holland, P.T. J. Agric. Food Chem. 41 388 (1993) 473 Berger, A. Chromatographia 41 133 (1995) 474 Garcia, F. and Henion, J. Journal of Chromatography 606 237–247 (1992) 475 Dinelli, G., Vicari, A. and Bonetti, A. Journal of Chromatography 700 195 (1995) 476 Raiser, R.W., Barefoot, A.C., Dietrich, R.F., Fogiel, A.J., Johnson, W.R. and Scott, M.T. Journal of Chromatography 554 91 (1991) 477 Howard, A.L. and Taylor, L.T. Journal of Chromatography Sci. 30 374 (1992) 478 Schneider, G.E., Koeppe, M.K., Naidu, M.V., Horne, P., Brown, A.M. and Mucha, C.F. J. Agric. Food Chem. 41 2404 (1993) 479 Nilve, G., Knutsson, M. and Jonsson, J.A. Journal of Chromatography 688 75 (1994) 480 Galletti, G.C., Bonetti, A. and Dinelli, G. Journal of Chromatography 692 27 (1995) 481 Cambon, J.P. and Bastide, J. J. Agric. Food. Chem. 41 333 (1996) 482 Volmer, D., Wilkes, J.G. and Leveson, K. Rapid Commun. Mass Spectrom. 9 767 (1995) 483 Di Corcia, A., Crescenzi, C., Samperi, R. and Scappaticcio, L. Analytical Chemistry 69 2819 (1997) 484 Martin-Esteban, A., Fernandez, P. and Camera, C. Analytical Chemistry 69 3267 (1997) 485 Zen, J.M., Jeng, S.H. and Chan, H.J. Analytical Chemistry 68 498 (1996) 486 Carpenter, R.A., Hollowell, R.H. and Hill, K.M. Analytical Chemistry 69 3314 (1997) 487 Ting-Long, Y., Guo, Y. and Qinhu, M. Analytical Chemistry 70 347 (1998)
Chapter 10
Miscellaneous organic compounds
10.1 Squoxin piscicide 10.1.1 Natural waters Kiigemagi et al. [1] developed a method for the determination of residues of this substance (1,1'-methylenedi-2-naphthol) in water. They used derivatization gas chromatography and spectrophotometric methods to determine down to 0.1ppm and to 2ppm, respectively, of this substance in water samples. Squoxin rapidly breaks down in water into a wide range of decomposition products. None of these products interferes sufficiently with the spectrophotometric method to hinder its usefulness for monitoring purposes. 10.2 Coprostanol 10.2.1 Natural waters and sewage effluents Recently, the use of coprostanol (5ß-cholestan–3ß-ol) as a molecular marker of faecal pollution of water has been suggested [2–5]. It has been shown that this saturated sterol satisfies the criteria for an indicator of faecal contamination of water [6]. Coprostanol satisfies the generally accepted criteria of a good indicator of faecal pollution. It is believed that the only source of this compound is the faeces of higher animals, including man. It is biodegradable and can be removed from domestic sewage by adequate treatment. Furthermore, it has been ambiguously proved that the concentration of coprostanol is highest in the overtly faecal polluted water and there is a progressive decrease in the concentration of this compound in the lesser polluted waters. Since its isolation and identification is unaffected by chlorination or by heat and toxic substances discharged from industrial outfalls the advantage of using a molecular rather than a biological indicator of faecal
Fluorescent whitening agents
629
pollution is further demonstrated. Such a characteristic of coprostanol is especially significant in the current trend promoting disinfection of raw and treated waste water. Because of this unique property, coprostanol might also be a useful indicator in monitoring the source, course and extent of faecal pollution in the ocean or brackish waters where bacteriological evidence is often doubtful. Dawson and Best [7] investigated the use of coprostanol as an internal tracer for faecal contamination in water by investigating the method of analysis and the effects of sewage treatment on the concentration of coprostanol, and by marine surveys at sewage outfalls. Analysis time is greater than for faecal bacteria but immediate analysis is not necessary and the results are less subject to error. Singley et al. [8] have described a gas chromatographic method for the analysis of coprostanol. This technique was used in extensive field studies, degradation studies and studies on treatment plant efficiency, and was also used as a standard for evaluating a colorimetric method that was developed and shown to be capable of determining coprostanol in polluted water at levels of 1µg L-1. It was shown that there was good correlation between coprostanol and biochemical oxygen demand (BOD), chemical oxygen demand (COD) and total organic carbon (TOC). Wun et al. [9] used XAD-2 resin for the analysis of coprostanol in river and lake water and secondary sewage treatment plants. They showed that extraction of coprostanol from water by adsorption on a column of Amberlite XAD-2 resin is as efficient as conventional liquid– liquid extraction. Maximal recovery depends on the pH value of the sample, flow rate, resin mesh size and concentration of the coprostanol. The final determination was carried out by gas chromatography. In further work, Wun et al. [10] improved the efficiency of the extraction of coprostanol using an XAD-2 resin column by decreasing the extraction time using a ‘closed’ column technique and by determining the effects of sample pH on adsorption processes. Coprostanol was strongly adsorbed to polystyrene XAD-2 adsorbents, at pH2, with 100% retention, and the adsorbed sterol was easily eluted with acetone adjusted to pH8.5–9.0 with ammonium hydroxide. It was also shown that with a closed column method, large volumes of water can be extracted in a relatively short time and with higher sensitivity than that of the liquid–liquid partitioning procedure. 10.3 Fluorescent whitening agents 10.3.1 Natural waters These substances can occur in river waters originating from their extensive use as ingredients of synthetic detergents for washing-up and
630
Miscellaneous organic compounds
cleaning operations. Abe and Yoshima [11] used thin-layer chromatography to study fluorescent whitening agents in river waters. Uchiyama [12] has given details of a procedure he developed for determining down to 0.2mg L-1 whitening agents and for the isolation and determination of fluorescent whitening agents in river water and bottom deposits. The fluorescent whitening agents were sodium salts of a sulphonated stilbene derivative and this was measured by fluorescence with the use of tetra-n-butyl ammonium hydroxide. Hellman [13] has used infrared spectroscopy to study the breakdown of minor ingredients of household detergent products, such as fabric whiteners and bleaching agents in river waters. Uchiyama [12] has discussed the separation and determination of fluorescent whitening agents in water. 10.4 Mestranol and ethynyloestradiol 10.4.1 Natural waters Okuno and Higgins [14] have described a procedure for determining residual levels of mestranol (17a-ethynyl-3-methoxyestra-1,3,5(10)-trien17ß-ol) animal damage control chemisterilant and its 3-hydroxy homologue, ethynyloestradiol in water samples. The lower limits of detection were 0.01 mg-1 for water samples. After extraction in acidic medium, samples are cleaned up by Florasil column chromatography. Water samples can then be analysed by gas–liquid chromatography. Thinlayer chromatography was used to confirm the results obtained by gas chromatography. Good recoveries were obtained from water samples. For a fieldcollected run-off sample spiked at 0.01ppm, recoveries averaged 99% for mestranol and 73% for ethynyloestradiol. The lower ethynyloestradiol recoveries in this case apparently reflect a less favourable partitioning of the compound into the chloroform phase. 10.5 Carboxymethyl succinate 10.5.1 Natural waters and sewage effluents A method has been described [15] for determining this detergent builder in river water and sewage effluent. This sample is centrifuged, adjusted to 1M in hydrochloric acid and, after 30min, recentrifuged. The supernatuant solution (1mL) is heated at 100°C for 30 min, then treated with 0.5% phenylhydrazine solution in 2M hydrochloric acid (0.3mL) and heated for a further 30min; 10% barium chloride solution (0.5mL), concentrated
Geosmin and other water flavouring chemicals
631
aqueous ammonia (0.15mL) and ethanol (6mL) are added and the mixture is kept at 4°C for 20h. The precipitate is centrifuged off and baked at 130°C for 30min, 0.05% 2-naphthol solution in 92.5% sulphuric acid (1mL) is added and the residue is completely dissolved by vigorous mixing. After heating the solution in a boiling water bath for 1h, 80% sulphuric acid (3mL) is added and the mixed solution is set aside at room temperature for 20min before the extinction is measured at 480nm against water. As little as 1µg of carboxymethyl succinate can be determined. 10.6 Isopropylmethylphosphonofluoridate (GB) and S-2(diisopropylamino)ethyl-Oethylmethylphosphonothioate (VX) 10.6.1 Sea water To determine these substances at the parts per trillion (US) level in sea water, Michel et al. [16] mixed the sample (0.1mL) with 0.1mL of eel cholinesterase solution in a buffer solution of pH7.2 (0.1M in morpholinopropane sulphonic acid, 0.01M in EDTA and containing 0.1% of gelatin). The mixture is incubated at 25°C for up to 30h. To determine residual enzyme activity, 0.1mL of substrate mixture (2nM in 5,5’dithiobis-(2-nitrobenzoic acid) is added and, after 1h, the extinction of the mixture is measured at 412nm. Standards are analysed similarly, and a blank determination is carried out without use of the enzyme. To determine VX alone, a 25mL sample of sea water is extracted with dichloromethane (2 × 5mL), and combined extracts are mixed with 0.1mL of 0.1M hydrochloric acid and the mixture is evaporated under nitrogen at room temperature. The residue is dissolved in water and aliquot is analysed as described above. 10.7 2,3-Dichloro-1,4-naphthaquinone 10.7.1 Natural waters A spectrophotometric method has been described for determining down to 2µg of this fungicide in water based on the formation of a coloured reaction product with aniline [17]. 10.8 Geosmin and other water flavouring chemicals Geosmin (trans 1,10-dimethyl-trans decanol) is a compound with a musty odour; it is a metabolite of some kinds of actinomycetes.
632
Miscellaneous organic compounds
10.8.1 Natural waters Persson [18] has discussed the determination of geosmin and 2methylisoborneol in water. Yasuhara and Fuwa [19] have described a quantitative method for determining geosmin in river water using computer-controlled mass fragmentography using a JOEL Model JMS-D 100 mass spectrometer connected with a JOEL Model JGC-20K gas chromatograph and a JOEL Model JMA-2000 mass data analysis system. The detection limit of geosmin was 10ng mL-1. 10.8.2 Potable waters 10.8.2.1 Gas chromatography–mass spectrometry Kikuchi et al. [20] identified geosmin in water supplies using gas chromatography–mass spectrometry. The water was chlorinated, treated with aluminium sulphate, and filtered through a layer of sand; it was then passed through a column of activated charcoal to adsorb odoriferous compounds. The charcoal was extracted several times with dichloromethane and the oily material recovered from the extract was distilled in water, the distillate was extracted with dichloromethane and the solvent was removed by evaporation. The residue was purified by chromatography on silica gel, with pentane and pentane– ethyl ether (99:1) as solvents. The eluate, which contained a musty smelling oil, was subjected to gas chromatography at 150°C on a stainless steel column (2m × 3mm) packed with 15% Reoplex 400 on Chromosorb WN AW with nitrogen as the carrier gas (30mL min-1) and flame ionization detection. The identity of the geosmin was confirmed by mass spectrometry and by further gas chromatography at 115°C on a glass column (1.5m × 2mm) packed with 25% PEG 21M Chromosorb W with helium as carrier gas (2kg cm-2). Otsuhara and Suwa [21] conducted studies on odorous compounds in reservoir water, filtration plant water and broth cultivated with actinomycetes and algae by gas chromatographic–mass spectrometric analysis. Odorous compounds were isolated from water by adsorption and extraction methods and identified as geosmin and 2methylisoborneol. Geosmin and 2-methylisoborneol were found in the surface water of a Japanese reservoir and Micromonospora isolated from a filtration plant produced 2-methylisoborneol. It was suggested algae were much concerned in the production of odorous compounds. Hwang et al. [22] have described a method for the determination in potable waters of geosmin and other earthy musty odorants (2isopropyl-3-methoxypyrazine, 2-isobutyl-3-methoxypyrazine, 2methylisoborneol and 2,3,6-trichloroanisole). These workers used
Ozonization products
633
closed-loop stripping coupled with gas chromatography and mass spectrometry to determine down to 0.8µg L-1 of these compounds. Increasing the ionic strength of the samples by adding sodium sulphate increased both the stripping rate and the sensitivity of the method. 10.8.2.2 Miscellaneous Krasner [23] has reviewed methods for the determination of geosmin in potable water. 10.9 Dihydroxyabietic acid 10.9.1 Trade effluents Brownlee and Strachan [24] determined dihydroxyabietic acid in receiving waters of a kraft pulp and paper mill. 10.10 Organic oxidants 10.10.1 Natural waters Larson et al. [25] have studied the occurrence and determination of organic oxidants in river water. They give details of methods to concentrate these compounds and determine them by iodiometry. 10.11 Ozonization products Since the detection of halogenated organics in potable water much research effort has been directed towards finding water treatment processes to remove such organics and their precursors and toward finding disinfectants other than chlorine. Great interest has been focused upon ozonization because both disinfection and organic removal can be accomplished with this process. As ozonated end-products will occur in water produced by such processes and these could be potentially toxic and would accumulate in waste water after repeated cycles of use it is necessary to ascertain what end-products occur in water that has been ozonated and subsequently chlorinated. 10.11.1 Natural waters Brunet et al. [26] studied the formation of organic materials during the ozonization of filtered surface water. They found that ozonization leads to a significant decrease in aromatic compounds together with an
634
Miscellaneous organic compounds
increase in polar compounds. Ozonization rates of the order 2mg L-1 appear to cause the degradation of the most reactive polyhydroxylated aromatic haloform precursors at pH7.0. There was an increase of precursors of the methyl ketone type which are very reactive with iodine and with chlorine at pH greater than 7. The results obtained by ultraviolet spectrometry and high-pressure liquid chromatography, were confirmed by data obtained by ozonization of dilute aqueous solutions of pure products. 10.11.2 Potable waters Glaze et al. [27] analysed ozonization byproducts produced before and after chlorination in potable water and waste water using size exclusion chromatography and halogen-specific microcoulometry. Gas chromatography with an electron-capture detector and gas chromatography– mass spectrometry was also used. Glaze et al. [27] showed that the majority of the organic halogen found in chlorinated lake water is not accounted for by trihalomethanes but is present, to a large extent as 2,4,6,2',4',6'-hexachlorobiphenyl. They studied the ozonization and photolytic ozonization of this compound. The products identified are shown in the proposed reaction scheme in Fig. 10.1. Except for the keto– enol pair shown in Fig. 10.1 all products, starting with 2-chloro-3-(2,4,6trichlorophenyl)maleic acid and beyond were identified from their mass spectra. Significantly, the scheme proposes that the ring first ruptured is degraded in a series of rapid steps before the second ring is attacked. The repetitive process which destroys these compounds is (1) double-bond cleavage resulting in aldehydes which (2) oxidize to carboxylic acids which then decarboxylate. Further cleavage naturally occurs when the system contains additional double bonds so that organic carbon is converted to carbon doxide via either intermediate carboxylic acids or the terminal compound in the ozone process, oxalic acid. Yamado and Somiya [28] have reported the quantitative determination of aldehydes and organic acids produced by ozonization of some model compounds in potable water. Aldehydes and ketones were determined by gas chromatography, oxoacids were converted to their methyl esters by diazomethane and then determined by gas chromatography, and dicarbonyl compounds were measured by a spectrophotometric method in alcoholic alkaline solution. Jolly et al. [29] have conducted a study into the non-volatile products produced during the chlorination, ozonization and ultraviolet irradiation of supply water and water produced in waste water treatment plants. High-performance liquid chromatography and mass spectrometry were used to separate organic constituents from concentrates of samples before and after treatment. Chromatographic
Fig. 10.1 Proposed reaction scheme for photolytic ozonolysis of hexachlorobiphenyl in aqueous solution. From CTMA (2-chloro-3-(2,4,6-trichlorophenyl)maleic acid) to end of scheme have been identified from mass spectra (unconfirmed by analysis of authentic sample). Reaction type code: C, Criegee cleavage; D, decarboxylation (RCO H-RH): R, replacement (RCL → ROH): (O) oxidation of aldehyde to carboxylic 2 acid; h, hydrolysis of acid chloride to acid; Short lines not terminated by a letter indicate chlorine atoms. (Reprinted with permission from Glaze et al. [27]. Copyright (1979) American Chemical Society.)
636
Miscellaneous organic compounds
profiles reveal the differences due to disinfection of the treated sample. High-performance liquid chromatographic analysis of chlorinated, ozonated and UV irradiated secondary effluents indicates that both chlorination and ozonization destroy chromatographic constituents and also produce chromatographic constituents. Ultraviolet irradiation at disinfection levels appears to have little chemical effect on the nonvolatile constituents separated by high-performance liquid chromatography. 10.11.3 Waste waters The ion-exclusion chromatographic procedure described in Section 10.11.2 [27] has also been applied to the analysis of waste waters. The high-performance liquid chromatographic procedure [29] described in Section 10.11.2 has also been applied to waste waters. Jolly et al. [30] characterized non-volatile organics produced in the ozonization and high chlorination of waste water. The treated effluents were analysed by high-performance liquid chromatography and gas– liquid chromatography as a means of detecting both nonvolatile and volatile organic constituents. While chlorinated primary effluent was repeatedly found to be mutagenic towards a particular strain, neither chlorinated, ozonated nor UV-irradiated secondary effluents exhibited mutagenic activity even when tested at 10–20-fold concentrations. 10.11.4 Sewage effluents Kuo et al. [31, 32] have studied the response of solutions of 2-propanol, acetic acid and oxalic acid to ozonization, with and without ultraviolet irradiation, in the laboratory. The effect of ultraviolet irradiation was to enhance the rate of oxidation of these organic compounds; the products of the reaction were identified by gas chromatography and the rate of decomposition determined in each case. Methylene chloride, chloroform, carbon tetrachloride and corresponding bromosubstituted derivatives were identified in the products of chlorination but not in those of ozonization, which appeared to convert the residual organic materials (humic and fulvic acids) to harmless products, alcohols, ketones, formaldehyde, lower fatty acids, oxalic acid, glyoxylic acid, pyruvic acid and ketomalonic acid, leading ultimately to the formation of carbon dioxide. Kuo et al. [31] examined the ozonization products gas chromatographically using a Hewlett Packard 5750B gas chromatograph equipped with dual flame ionization detectors. The derivative gas chromatographic procedures of Bethge and Lindstrom’s method [33]
Ozonization products
637
were followed to determine formic, acetic, and propionic acids as their benzyl esters. Oxalic acid was determined gas chromatographically after the compound was converted to methyl ester using diazomethane [34]. Glyoxylic acid was determined by a colorimetric method described by Kramer et al. [35]. The chromatropic acid method was used to determine formaldehyde [36]. 2-Propanol was oxidized by ozonization and ultraviolet irradiation to acetone, which in turn was oxidized to acetic and oxalic acids. Trace amounts of formaldehyde and formic acid were also detected in the ozonated acetone mixtures. Pyruvic and ketomalonic acids may be among the undetermined ozonization products. Ozonization of acetic acid resulted in the formation of glyoxylic acid. Glyoxylic acid was, however, present in trace amounts (98%) recoveries of inorganic arsenic (III) and of the three organic arsenic compounds mentioned above. Sandhu and Nelson [1] have also studied the interference effects of several metals on the determination of organically bound arsenic at the 0– 100µg L-1 range in waste water by the silver diethyldithiocarbamate method. Antimony and mercury interfere specifically, forming complexes with silver diethyldithiocarbamate at absorbance maxima at 510 and 425nm respectively. Recovery of arsenic released by digesting solutions was tested and shown to give about 90% recovery of organic arsenic. 12.2 Organolead compounds 12.2.1 Natural waters 12.2.1.1 Gas chromatography Chau et al. [36] have described a simple and rapid extraction procedure to extract the five tetraalkyllead compounds (Me Pb, Me Et Pb, MeEt Pb, 4
3
2
3
730
Organometallic compounds
Me Et Pb and Et Pb) from natural water. The extracted compounds are 2 2 4 analysed in their authentic forms by a gas chromatographic atomic absorption spectrometry system. Other forms of inorganic lead do not interfere. The detection limit for water (200mL) is 0.50µg L-1. The main interest of Chau et al. [36] was in the determination of organically bound lead produced by biological methylation of inorganic and organic lead compounds in the aquatic environment by microorganisms. The gas chromatographic–atomic absorption system used by Chau et al. (used without a sample injection trap) for this procedure has been described [37]. The extract was injected directly into the column injection port of the chromatograph. Instrumental parameters were identical. A Perkin–Elmer electrodeless discharge lead lamp was used; peak areas were integrated. Potter et al. [38] have applied gas chromatography and thin-layer chromatography to the detection and determination of alkyllead compounds and alkyllead salts in natural waters. The limit of detection of this method was 0.5–1.0µg alkyllead salt. The total concentration of lead in natural waters is generally low owing to absorption into sediment unless the pH is exceptionally low. This explains the observation that the levels of lead in rain water are twice those in ground water. Tetraalkyllead is insoluble in, and denser than, water and would be expected to accumulate with the sediment. Alkyllead salts are generally much more soluble in water than the corresponding organic lead salts and would remain in solution in the absence of suspended solids. However, alkyllead salts at low concentrations are totally absorbed into a variety of natural sediments which are nearly always present in natural waters. Chau et al. [39, 40] developed a gas chromatographic method for the determination of ionic organolead species and applied it to samples of lake Ontario water. When a 1L of sample water is taken, the following species can be determined in amounts down to 0.1µg L-1: Me2Pb2+, Me Pb+, Et Pb2+, Et Pb+ and Pb2+. 2 2 3 The highly polar ions are quantitatively extracted into benzene from aqueous solution after chelation with dithiocarbamate. The lead species are butylated by Grignard reagent to the tetraalkyl form, R PbBu (R = n (4–n) CH or C H ) and Bu Pb, all of which can be quantified by a gas 3 2 5 4 chromatography–atomic absorption spectrometry method. Molecular covalent tetraalkyllead species, if present in the sample, are also extracted and quantified simultaneously. Other metals coextracted by the chelating agent do not interfere. All the ionic alkyllead compounds slowly degrade in the presence of light. However, lake water samples enriched with dimethyllead chloride and trimethyllead acetate in 100µg L-1 level are stable over a period of at least one month in the laboratory when stored in the dark
Organolead compounds
731
and refrigerated. There is no need to add any preservative to the sample. Storage in a cold dark room is recommended. Alternatively, the samples can be extracted, butylated and dried over anhydrous sodium sulphate. Tetraalkyllead compounds (typically tetramethyllead, trimethyllead, dimethyldiethyllead, methyltriethyllead and tetraethyllead) have high vapour pressures and are seldom found in water unless they are adsorbed on sediments or particulate matter. If these compounds are present in the water sample, they can be extracted into benzene and included in the determination. The butylation reaction has no effect on the tetralkyllead. Chakraborti et al. [41] analysed water samples for dialkyl and trialkyllead compounds by gas chromatography–atomic absorption spectrometry. Analysis of 500mL of water enables the determination of 1.25mg L-1 lead for PbMe + and 2.5ng L-1 for PbEt +. Extraction recoveries 3 2 are in excess of 90% for all compounds studied. Rapsomanikis et al. [42] speciated lead and methyllead ions in water by gas chromatography–atomic absorption spectrometry after elution with sodium tetraethylborate. This purge and trap technique has detection limits of 0.2ng L1-1 for trimethyllead and trimethyllead species on 50mL water samples. 12.2.1.2 Polarography Colombini et al. [44] have described a technique for the determination of organometallic species in natural waters based on the selective organic phase extraction coupled with differential pulse polarography. The analytical procedure was applied to alkyllead compound speciation and found to be reliable for the individual detection and determination of organolead complexes at trace levels in natural waters including sea water. 12.2.1.3 Atomic asborption spectrometry De Jonghe et al. [43] have described an extraction technique for determining traces of trialkyllead compounds in water samples, based on the salting out of the trialkyllead ions as neutral species into an inorganic solvent, followed by graphite furnace atomic absorption spectrometry. A detection limit of 0.02µg L-1 was achieved with samples. 12.2.1.4 Miscellaneous Jarvie et al. [45] studied the reactions between trimethyllead chloride and sulphides in aqueous systems. Factors affecting these reactions have been
732
Organometallic compounds
investigated. It is concluded that the formation of tetramethyllead from trimethyllead compounds in natural waters is due to chemical reactions, and not to biomethylation. The number of analytical techniques that can be used for lead analysis in water is considerable. However, at the concentration levels found in environmental samples the choice is much more limited. Only three techniques enable sub-ppb detection limits, atomic absorption spectrometry, anodic-stripping voltammetry and inductively coupled plasma–mass spectrometry [219]. The cost of atomic absorption spectrometry instrumentation is relatively high and that of inductively coupled plasma–mass spectrometry is prohibitive. None of these techniques can be easily adopted to field measurements. Anodic-stripping voltammetry instrumentation can be portable, yet natural water samples must usually be mineralized prior to final determination due to adverse matrix effects caused by organic compounds, a task that is not easily accomplished in the field. 12.2.1.5 Preconcentration Mikac and Branica [46] preconcentrated dissolved dialkyllead and inorganic lead species by coprecipitation with barium sulphate. Alkyllead was then determined in the concentrate by differential pulse anodicstripping voltammetry. 12.2.2 Rain water 12.2.2.1 High-performance liquid chromatography Blaszkewicz el al. [47] determined trialkyllead species in rain water. Interfering metal ions in rain water were complexed with EDTA before adjustment of the pH to 10. Samples were pumped through an extraction column of silica gel to absorb lead compounds which were then desorbed with acetate buffer containing methanol at pH3.7. The eluate was diluted and adjusted to pH8 with borate buffer before further concentration on a Nucleosil 10–C precolumn and separated 18 with methanolic acetate buffer. Online detection used a postcolumn chemical reaction detector. Detection limits for sample volumes of 500mL were 15pg mL-1 and 20pg mL-1 for trimethyl and triethyllead respectively. Standard deviation was less than 4% for a sample containing 90pg triethyllead per millilitre. Concentrations of trimethyland triethyllead in rain, melt water and surface water samples were between 20 and 100pg mL-1.
Organolead compounds
733
12.2.3 Waste waters 12.2.3.1 Spectrophotometric method Imura et al. [48] have described a Spectrophotometric procedure employing 1-hydroxy-4-(4-nitrophenylazo)-2-naphthoate as chromogenic reagent for the determination of triethyllead ions in industrial waste waters. The coloured adduct is extracted with chloroform. The absorption maximum is at 440nm and the optimum pH for the extraction from 1% aqueous sodium chloride is 8.1–8.3. In the determination of about 60µg of triethyllead ions, dimethyl- and diethyllead ions (100µg) and Pb2+ (1.8mg) are masked with 0.01M ethylenedediamine-N,N’-bis-(2hydroxyphenylacetic acid) (disodium salt) (2mL) and copper and ferrous iron are masked with 0.01M 1,2-diaminocyclohexane-N,N,N’,N’tetraacetate (2mL). Several other ions do not interfere. The limit of determination is 0.2ppm of triethyllead. 12.2.3.2 Atomic absorption spectrometry Aneva [49] has discussed the determination of traces of tetraalkyllead compounds in waste waters by graphite furnace absorption spectrometry. Tetraalkyllead compounds were extracted from water with hexane, converted to water soluble iodides by reaction with iodine in the extract and re-extracted in dilute nitric acid. Determination was by graphite furnace atomic absorption spectrometry against an inorganic lead standard. The mean recovery of lead was 95% and no interference from other metal and non-metal ions was experienced. The method could be used for samples containing down to 5µg L-1 lead. 12.2.4 Trade effluents 12.2.4.1 Spectrophotometric method The method [48] described in Section 12.2.3.1 for the determination of triethyllead ions in waste waters has also been applied to trade effluents. 12.2.5 Potable waters 12.2.5.1 Gas chromatography–mass spectrometry Gorecki and Pawliszyn [218] have described a new gas chromatographic method for the determination of tetraethyllead and ionic lead in water by solid-phase microextraction gas chromatography. Tetraethyllead is
734
Organometallic compounds
extracted from the headspace over the sample. Inorganic lead is first derivatized with sodium tetraethylborate to form tetraethyllead, which is extracted in the same way as pure tetraethyllead samples. The analytical procedure was optimized with respect to pH, amount of derivatizing reagent added, stirring conditions, and extraction time. The detection limit obtained for tetraethyllead was found to be 100 ppt when using a flame ionization detector and 5ppt when using an ion-trap mass spectrometer. The detection limit for Pb2+, limited by the non-zero blank, was found to be 200ppt. Linear calibration curves were obtained for both analytes when a flame ionization detector was used for detection. For lead they spanned over four orders of magnitude. The ion-trap mass spectrometry offered excellent sensitivity and selectivity, but the calibration curves were non-linear when the m/z = 295 ion was used for quantitation. The method has been verified on spiked tap water samples. An excellent agreement was found between the results obtained for standard solutions prepared using NANO pure water and spiked tap water samples. 12.3 Organomercury compounds 12.3.1 Natural waters The two main techniques for the determination of mercury in water samples are atomic absorption spectrophotometry and gas chromatography. Mercury in water samples can exist in inorganic or organic form, or both. Normal methods of reducing inorganic mercury compounds to mercury with reagents such as stannous chloride do not work with organomercury compounds and hence organomercury compounds are not included in such determinations. Owing to the conversion of Hg2+ to CH Hg+ in river waters, and to the presence of 3 mercury in a large number of organic pollutants, it is often observed that a high percentage of the mercury is present in the form of organic compounds. Some organic mercurials like methylmercury chloride and dimethylmercury may be reduced by a combination of cadmous chloride, but this method requires large quantities of reductants and the use of strong acid and strong alkali [50]. Organic mercury compounds can be decomposed by heating with strong oxidizing agents such as potassium chromate or nitric acid– perchloric acid, followed by reduction of the formed divalent mercury to mercury vapour [51, 52]. Both methods are rather time-consuming and not very suitable for automation. Potassium persulphate has also been used to aid the oxidation of organomercury to inorganic mercury compounds and this forms the basis of an automated method [53].
Organomercury compounds
735
Goulden and Afghan [54] have used ultraviolet irradiation as a means of decomposition, following the original proposal of Armstrong et al. [55]. After the photochemical oxidation, the formed inorganic mercury is reduced to metallic mercury in the usual way by stannous chloride. This method reduces the consumption by oxidizing agents and thus diminishes considerably the risk of contamination; it also leads to shorter analysis times. Determination with and without irradiation enable the separate determination of total and inorganic mercury respectively. Bennett et al. [56] later showed that acid permanganate alone did not recover three methyl mercuric compounds, while the addition of a potassium persulphate oxidation step increased recoveries to 100%. El-Awady et al. [57] confirmed the low recoveries of methylmercury by acid permanganate. They showed that only about 3% of methylmercury could be recovered by this method, while the use of potassium persulphate produced complete recovery. 12.3.1.1 Gas chromatography Nishi and Horimoto [58, 59] determined trace amounts of methyl-, ethyl, and phenylmercury compounds in river waters and industrial effluents. In this procedure, the organomercury compound present at less that 0.4ng L-1 in the sample (100–500mL) is extracted with benzene (2 × 0.5 vol. relative to that of the aqueous solution). The benzene layer is then backextracted with 0.1% L-cysteine solution (5mL), and recovered from the complex by extracting with benzene (1mL) in the presence of hydrochloric acid (2mL) and submitted to gas chromatography using a stainless steel column (197cm × 3mm) packed with 5% of diethylene glycol succinate on Chromosorb W (60–80 mesh) with nitrogen as carrier gas (60mL min-1) and an electron-capture detector. The calibration graph is rectilinear for less than 0.1µg of mercury compound per millilitre of the cysteine solution. This method is capable of determining mercury down to 0.4µg L-1 for the methyl and ethyl derivatives and 0.86µg L-1 for the phenyl derivative. The above method has been modified [59] for the determination of methylmercury (II) compounds in aqueous media containing sulphur compounds that affect the extractions of mercury. The modified method is capable of handling samples containing up to 100mg of various organic and inorganic sulphur compounds per 100mL. The aqueous test solution (150mL) containing 100mg of methylmercury ions per 100mL is treated with hydrochloric acid until the acid concentration is 0.4%, then 0.3–1g of mercuric chloride is added (to displace methyl mercury groups bonded to sulphur), and the mixture is filtered. The filtrate is treated with aqueous ammonia in excess to precipitate the unconsumed inorganic mercury which is filtered off; this filtrate is made 0.4% in
736
Organometallic compounds
hydrochloric acid and extracted with benzene. The benzene solution is shaken with 0.1% L-cysteine solution, the aqueous phase is acidified with concentrated hydrochloric acid and then shaken with benzene for 5min and this benzene solution is analysed by a gas chromatography as described above. Another application of gas chromatography to natural water analysis is that of Longbottom [60] who used a Coleman 50 mercury analyser system as a detector. A mixture of dimethyl-, dipropyl- and dibutylmercury (1mg of each) was separated on a 6ft (1.8m) column packed with 5% DC-200 and 3% QF-1 on Gas Chrom Q and temperature programmed from 60 to 180°C at 20°C min-1. The mercury detector system was used after the column effluent had passed through a flame ionization detector; the heights of the resulting four peaks were related to the percentage of mercury in the compounds. Dressman [61] also used the Coleman 50 system in his determination of dialkylmercury compounds in river waters. These compounds were separated in a glass column (1.86m × 2mm) packed with 5% of DC-200 plus 3% of QF-1 on Gas Chrom Q (80–100 mesh) and temperature programmed from 70 to 180°C at 20°C min-1, with nitrogen as carrier gas (50mL min-1). The mercury compound eluted from the column was burnt in a flame ionization detector, and the resulting free mercury was detected by a Coleman mercury analyser MAS-50 connected to the exit of the flame ionization instrument; down to 0.1mg of mercury could be detected. River water (1L) was extracted with pentane–ethyl ether (4:1) (2 × 60mL). The extract was dried over sodium sulphate, evaporated to 5mL and analysed as above. Ealy et al. [62] have discussed the determination of methyl-, ethyl-and methoxymercury (II) halides in water. The mercury compounds were separated from the sample by leaching with 1M sodium iodide for 24h and then the alkylmercury iodides were extracted into benzene. These iodides were then determined by gas chromatography with 5% cyclohexanesuccinate on Anakron ABS (70–80 mesh) and operated at 299°C with nitrogen (56mL min-1) as carrier gas and electron-capture detection. Good separation of chromatographic peaks was obtained for the mercury compounds as either chlorides, bromides or iodides. The extraction recoveries were monitored by the use of alkylmercury compounds labelled with 208Hg. Zarnegar and Mushak [63] have described a gas chromatographic procedure for the determination of organomercury compounds and inorganic mercury in natural water. The sample is treated with an alkylating or arylating reagent and the organomercury chloride is extracted into benzene. Gas chromatography is carried out using electron-capture detection. The best alkylating or arylating reagents were pentacyano(methyl)cobalt (III) and tetraphenylborate. Inorganic and
Organomercury compounds
737
organic mercury could be determined sequentially by extracting and analysing two aliquots of sample, of which only one had been treated with alkylating reagent. The limits of detection achieved in the method were 10–20ng. Cappon and Crispin Smith [64] have described a method for the extraction, clean-up and gas chromatographic determination of organic (alkyl and aryl) and inorganic mercury in water. Methyl-, ethyl- and phenylmercury are first extracted as the chloride derivatives. Inorganic mercury is then isolated as methylmercury upon reaction with tetramethyltin. The initial extracts are subjected to thiosulphate clean-up and the organomercury species are isolated as the bromide derivatives. Total mercury recovery ranges between 75 and 90% for both forms of mercury, and is assessed by using appropriate 203Hg-labelled compounds for liquid scintillation spectrometric assay. Specific gas chromatographic conditions allow detection of mercury concentrations of 1µg L-1 or lower. Mean deviation and relative accuracy average 3.2 and 2.2% respectively. Bowles and Apte [220] have described a method for the determination of methylmercury compounds in natural waters using steam distillation followed by gas chromatography with an atomic fluorescence spectrometric detector. These workers evaluated steam distillation as a technique for the separation of methylmercury compounds from water and obtained recoveries in spiking experiments ranging from approximately 100% in fresh waters and estuaries to 80% in sea water. The addition of ammonium pyrrolidine dithiocarbamate improved recoveries from sea water to 85%. The codistillation of inorganic mercury was prevented by the addition of ammonium pyrrolidine dithiocarbamate. The precision achieved in this method on Milli-Q water was 0.2% r.s.d. at the 0.2ng L-1 methylmercuric chloride level and 1.6% r.s.d. at the 2ng L-1 level. A detection limit of 0.24ng L-1 was achieved for a 50mL water sample. 12.3.1.2 Atomic absorption spectrometry Kalb [65] used concentrated nitric acid to decompose organomercury compounds in water samples prior to estimation by flameless atomic absorption spectroscopy. Stannous chloride is used to liberate elementary mercury, which is then vaporized by passing a stream of air (1360mL min1 ) through the solution. The air stream passes over silver foil, where mercury is retained by amalgamation and other volatile substances pass out of the system. The foil is heated at 350°C in an induction coil, and the air stream carries the mercury vapour through a cell with quartz windows. The atomic absorption at 253.65nm is measured, and the
738
Organometallic compounds
mercury concentration (up to 0.2ppm) is determined by reference to a calibration graph. Umezaki and Iwamoto [66] differentiated between organic and inorganic mercury in river samples. They used the reduction–aeration technique described by Kimura and Miller [67]. By using stannous chloride in hydrochloric acid only inorganic mercury is reduced, whereas stannous chloride in sodium hydroxide medium in the presence of cupric copper reduces both organic and inorganic mercury. The mercury vapour is measured conventionally at 254nm. Ions that form insoluble salts or stable complexes with Hg(II) interfere. Doherty and Dorsett [68] analysed environmental water samples by separating the total organic and inorganic mercury by electrodeposition for 60-90min on a copper coil in 0.1M nitric acid medium and then determined it directly by flameless atomic absorption spectrophotometry [69, 70]. The precision and accuracy are within ± 10% for the range 0.1– 10 parts per 109. The sensitivity is 0.1 part per 109 (50mL sample). A tentative method has been described for the determination of mercury in water [71]. This method determines all forms of mercury provided that they are first converted to inorganic mercury. All forms of mercury in non-saline waters are converted to inorganic mercury using a prolonged oxidation with potassium permanganate. The inorganic mercury is determined by the flameless atomic absorption spectrophotometric technique using a method similar to that described by Osland [72]. Acid stannous chloride is added to the sample to produce elemental mercury: Hg2+ + Sn2 → Hg0 + Sn4+ The mercury vapour is carried by a stream of air or oxygen into a gas cuvette placed in the path of the radiation from a mercury hollowcathode lamp and the absorption of this radiation at 253.7nm by the mercury vapour is measured. Many of the potential interferences in the atomic absorption procedure are removed by the preliminary digestion– oxidation procedure. The most significant group of interfering substances is volatile organic compounds which absorb radiation in the ultraviolet. Most of these are removed by the pretreatment procedure used and the effect of any that remain is overcome by pre-aeration. Bromide and iodide ions may cause interference. Substances which are reduced to the elemental state by stannous chloride and then form a stable compound with mercury may cause interference; e.g. selenium, gold, palladium and platinum. The effects of various anions, including bromide and iodide, were studied. These are not likely to be important interferers. Excellent performance characteristics are presented for this method (Table 12.2). Sampling techniques are described in detail,
Organomercury compounds
739
Table 12.2 Performance characteristics, Water Research Centre method for mercury
including methods of cleaning sample bottles and fixing the sample with a solution of potassium dichromate in nitric acid. Farey et al. [73] have discussed ultraviolet photochemical systems for the decomposition of organomercury compounds prior to analysis by cold-vapour atomic fluorescence spectroscopy. These workers compared the effectiveness of a bromination treatment for the liberation of mercury from organomercury compounds with a pretreatment procedure involving oxidation with a permanganate– sulphuric acid mixture recommended by workers at the Water Research Centre, UK [74]. The basis of the bromination technique, in which a bromate– bromide reagent in hydrochloric acid reagent is used to generate bromine in the sample, is that the bromine quantitatively cleaves both alkyl- and arylmercury compounds to inorganic mercury bromide. Recoveries of inorganic mercury from distilled water spiked with phenylmercury (II), chloride, thiomersal, ethylmercury (II) chloride, methylmercury (II) chloride, phenylmercury (II) acetate and ptolymercury (II) chloride are greater than 95%. Standard deviation is 0.1– 0.2µg L-1 at the 1–2µg L-1 level. Farey et al. [73] claim that their treatment compares favourably with an established permanganate–sulphuric acid method. An advantage of the technique is that it can easily be carried out while sampling on-site. The sample is collected in glass bottles containing hydrochloric acid and the bromate–bromide solution is added. A bromination reaction time is
740
Organometallic compounds
then provided from the collection of the sample to the analysis in the laboratory and this is far in excess of that necessary to decompose the organic mercury. In addition, as aqueous mercury (II) solutions are stabilized by strong oxidizing agents, the oxidizing conditions so created will help to preserve the inorganic mercury formed. Goulden and Afghan [54] have used ultraviolet irradiation as a means of decomposition of organomercury compounds, following the original proposal of Armstrong et al. [55]. After the photochemical oxidation, the formed inorganic mercury is reduced to metallic mercury in the usual way using stannous chloride. The method reduces the consumption by oxidizing agents and thus diminishes considerably the risk of contamination; it also leads to shorter analysis times. Determination with and without irradiation enable the separate determination of total and inorganic mercury respectively. Bennett et al. [56] showed that acid permanganate alone did not recover three methylmercuric compounds while the addition of potassium persulphate oxidation step increased recoveries to 100%. El-Awady et al. [88] confirmed the low recoveries of methylmercury by acid permanganate. They showed that only about 3% of methylmercury could be recovered by this method while the use of potassium persulphate produced complete recovery. Kiemeneij and Kloosterboer [89] have described an improvement on the Goulden and Afghan photochemical decomposition of organomercury compounds in the microgram per litre range in water prior to determination by cold-vapour atomic absorption spectrophotometry. Decomposition of the organomercurials is carried out by means of ultraviolet radiation of a suitable wavelength from small, low-pressure lamps containing in their cathodes either zinc, cadmium or mercury, or a mixture of these metals. The formed inorganic mercury is determined in the usual way by cold-vapour atomic absorption after reduction of divalent mercury to mercury vapour. Determinations with and without irradiation make possible separate determinations of total and inorganic mercury, respectively, in about 20min. Abo-Rady [75] has described a method for the determination of total inorganic plus organic mercury in nanogram quantities in water. This method is based on the decomposition of organic and inorganic mercury compounds with acid permanganate, removal of excess permanganate with hydroxylamine hydrochloride, reduction to metallic mercury with tin and hydrochloric acid, and transfer of the liberated mercury in a stream of air to the spectrometer. Mercury was determined by using a closed, recirculating air stream. Sensitivity and reproducibility of the ‘closed system’ was better, it is
Organomercury compounds
741
claimed, than those of the ‘open system’. The coefficient of variation was 13.7%. Goulden and Anthony [76] describe how the sensitivity of an automated cold-vapour atomic absorption method for mercury can be improved by equilibrating the reduced sample with a small volume of air at 90°C, to achieve a detection limit of 1ng mercury. Inorganic mercury, arylmercury compounds and alkylmercury compounds can be distinguished by changing the chemical reduction system. The authors suggest that their method of chemical speciation can be applied to natural waters. Other workers who have made earlier contributions to the determination of organically bound mercury compounds and inorganic mercury compounds by flameless atomic absorption spectroscopy include Baltisberger and Knudson [77], Bisagni and Lawrence [78], Frimmel and Winckler [79], Chan and Saitoh [81], Umezaki and Iwamoto [66], Stainton [82], Carr at al. [83], Fitzgerald et al. [84], Watling and Watling [80], Graf et al. [85], Simpson and Nickless [86] and Lutze [87]. 12.3.1.3 Inductively coupled plasma mass spectrometry Tong et al. [90] have shown that direct-injection nebulization– inductively coupled plasma mass spectrometry is a feasible choice for the automated determination of mercury and organomercury. 12.3.1.4 Photochroism-induced photoacoustic spectrometry This technique is applicable to the determination of down to 3ppt of mercury as its dithizonate in water samples [91]. 12.3.1.5 Neutron activation analysis Becknell et al. [92] first converted the organomercury to mercuric acid using chlorine. The mercury was then concentrated by removal as HgCl 24 by passing the sample solution (500mL, adjusted to be 0.1M in hydrochloric acid) through a paper filter disc loaded with SB-2 ionexchange resin. The paper was then heat sealed in Mylar bags and irradiated for 2h in a thermal neutron flux of ~1.3 × 1013 neutrons cm-2 s-1, after which the concentration of mercury in the sample was determined by the comparison method using a 77keV γ-ray photopeak from the decay of 197Hg.
742
Organometallic compounds
12.3.1.6 Miscellaneous Radiochromatographic assay has been used [93] as the basis of a method for the determination of inorganic mercury and methylmercury in river water. Schintu et al. [128] preconcentrated organic and inorganic mercury by extraction with dithizone solution. Organic mercury was then backextracted with aqueous sodium thiosulphate prior to determination by cold-trap cold-vapour atomic absorption spectrometry. Fitzgerald et al. [84] reported a cold-trap preconcentration technique for the determination of trace amounts of mercury in water. Kramer and Neidhart [129] determined ppb (µg L-1) levels of mercury by using aniline–sulphur resin for selective enrichment of mercury from surface waters. Chan and Saitoh [81] reported a method for the determination of submicrogram amounts of mercury in lake water based on dithizone extraction. Preconcentration of mercury prior to the measurement has also been achieved by amalgamation with noble metals [130–133]. Ke and Thibert [94] have described a kinetic microdetermination of down to 0.05µg mL-1 of inorganic and organic mercury in river water and sea water. Mercury is determined by use of the iodide-catalysed reaction between Ce(IV) and As(III), which is followed spectrophotometrically at 273nm. Van Ettekoven [95] has described a direct semiautomated scheme based on ultraviolet light absorption for the determination of total mercury in water and sewage sludge. The full determination time is about 10min. The lower limit of detection of mercury in water is 0.03µg L-1 and 0.2ppm (mg kg-1 dry matter) in sewage sludge. Matsunaga et al. [96] have discussed possible errors caused prior to measurement of mercury in natural water and sea water. Evans and McKee [97] give details of equipment and procedure for the determination of divalent mercury and organomercury compounds in amounts down to 1–2µg L-1 by electrochemical detection in the reductive mode at a gold–amalgamated mercury electrode combined with reversed-phase liquid chromatography. Potentiometric titration with standard solutions of dithiooxamide at pH5–6 has been used to estimate less than 100µg mercury in water samples [98]. The precision in the range 0.05–1.0µg L-1 mercury is about 4%. The first derivative should be used for end-point determination. A wide variety of ions can be tolerated but silver, copper and chloride interfere, and must be separated in a preliminary step. Braun et al. [99] showed that polyurethane foam loaded with diethyldithiocarbamate is suitable for the concentration of trace amounts of organic and inorganic mercury from natural water samples prior to chemical analysis. Mercury (II), methylmercury and phenylmercury ions
Organomercury compounds
743
at around 1µg L-1 levels can be almost quantitatively preconcentrated from water and eluted from the foam with acetone. Glockling [100] has studied the degradation of organomercury compounds in water. Kudo et al. [101] measured the concentrations of methylmercury and total mercury in Canadian and Japanese rivers. In all cases methylmercury was found to represent about 30% of the total mercury. Braun et al. [102] used radioisotope-induced X-ray fluorescence spectrometry to determine phenylmercury, methylmercury and inorganic mercury in river water after preconcentration on diethylammonium diethyldithiocarbamate-loaded polyurethane foam. Ahmed and Stoeppler [103] carried out decomposition and stability studies of methylmercury in river water using atomic cold-vapour absorption spectrometry. The behaviour was studied of methylmercuric chloride in water when exposed to light (irradiation samples with a 150W mercury lamp from a distance) and heat in the presence of different acids. Storage of mercury-containing samples The problems of preserving mercury in solution are well known. Although controversy still exists over which preservative is best, agreement on several of the factors which affect the stability of mercury solutions seems to have been reached. For example, it is agreed that low pH values, high ionic strengths and oxidizing environments help in keeping mercury in solution. Acids such as sulphuric acid [104], nitric acid [105-111], and hydrochloric acid [112] have been widely used in different amounts. Oxidants such as permanganate [57, 113–120] and dichromate [57, 118–120] have been shown to prevent volatilization of mercury. Sodium chloride and gold [119, 120] have also been used as preservatives. Various workers have commented on the instability of mercury solutions when stored in polyethylene or polypropylene containers [109, 111, 121, 122]. Carron and Agemian [123] have pointed out that whilst the majority of fresh water samples rarely contain mercury at levels over 0.5µg L-1 and in most cases 0.2µg L-1, most previous investigators of the stability of mercury solutions have carried out their tests at higher mercury levels. These workers studied the preservation methods which provide both low pH values as well as oxidizing environments using both synthetic and natural samples in a variety of containers, in order to obtain a practical method which would be adaptable to routine analysis for mercury in natural waters at sub-microgram per litre levels by the automated cold-vapour atomic absorption technique. The essential requirement was that the preservation method should maintain mercury
744
Organometallic compounds
in waters of low salt content (low conductivity such as distilled water) and of high salt content (high conductivity). The outcome of this work was that Carron and Agemian [123] recommended glass containers washed with concentrated nitric acid or chromic acid and a preservative consisting of a mixture of 1% sulphuric acid and 0.05% potassium dichromate. This preservative gives good accuracy, precision and low detection limits. It was also observed that the presence of methylmercury ions improves preservation efficiency. Stoeppler and Matthes [124] have studied the storage behaviour of methylmercury chloride and inorganic mercury in sea water samples. Carr and Wilkniss [106] showed that if a water sample is acidified to pH1 with nitric acid, then of any mercury that is initially associated with particulate matter in the sample about 80% enters solution during storage for a period of about 1 week. There is no significant loss to the container. If the sample is not acidified, most of the mercury is retained by the particulate matter, 15% is adsorbed by the container and only 10% remains in solution. 12.3.1.7 Preconcentration Vinyl dithiocarbamate resins Minagawa et al. [125] have described a technique employing chelating resins which has been applied to the determination of very low concentrations down to 0.2ng L-1 of organic and inorganic mercury in natural waters including rivers, lakes and rainwaters. The resin used contains dithiocarbamate groups which bind mercury but not alkali and alkaline earth metals. Both forms of mercury are collected at pH1-11 and eluted with slightly acid 4% thiourea in water. Large volumes of water can be concentrated to determine mercury by cold-vapour atomic absorption spectrometry. Mercury vapour is generated from inorganic mercury with alkaline stannous chloride and from organic and inorganic mercury with a cadmium chloride–stannous chloride solution. No interference was produced in the determination of 0.1µg of mercury (II) by the presence of at least 1000µg of each of the following ions or substances added to 5L aliquots of river water: chromium (III), magnesium, sodium, potassium, calcium, nickel (II), copper (II), lead (II), cadmium, gold (III), iron (III), aluminium, zinc, phosphate, chloride, carbonate, nitrate, sulphate, silicate, cysteine and humic acid. The accuracy of the method was tested by analysing river water samples spiked with known amounts of mercury and CH Hg+. The accuracy of 3 the method was satisfactory. The precision of the method was 9.5 ±
Organomercury compounds
745
0.43ng L-1 at the 10ng L-1 level and 0.36ng L-1 at the 15ng L-1 level for inorganic and organic mercury respectively. Yamagami et al. [126] also applied the chelating resins (dithiocarbamate type) to the determination of micrograms per litre of mercury in water. The samples are adjusted to pH2.3 and passed through a column packed with 5g of the resin, at a flow rate of 50mL min-1. The resin is then digested under reflux with concentrated nitric acid and the mercury is determined by atomic absorption spectrophotometry, using the reduction aeration technique. The method is relatively simple and inexpensive and the detection limit is 10ng mercury in water samples as large as 10L. Sulphydryl cotton f ibre Lee [127] used sulphydryl cotton fibre to preconcentrate methyl and ethylmercury compounds from natural waters. Adsorbed mercury compounds were desorbed from the fibre with hydrochloric acid– sodium chloride and then extracted into benzene prior to determination by electron-capture gas chromatography. In further work, Lee and Mowrer [222] preconcentrated methylmercury from natural waters on to a sulphydryl cotton fibre adsorbent using the column technique or the batch-column two-stage technique. A small volume of 2M hydrochloric acid was used to elute methylmercury and to separate it from inorganic mercury; 0.4–0.6 mL of benzene was used to extract methylmercury from the eluate. Analysis was performed by capillary gas chromatography with electron-capture detection. The detection limit for methylmercury was 14% of total S). Reduced non-sulphate inorganic sulphur compounds were not detected in pond water samples. Sulphur has been determined in water by methods based on the use of a cadmium sulphide membrane electrode [97], a silver–silver chloride electrode [98] and by microcoloulometry [99]. The cool-flame chemiluminescence emission spectrometry technique [226] described in Section 13.6.1.5 for the determination of phosphorus has also been applied to the determination of sulphur. 13.8.2 Waste waters 13.8.2.1 Emission spectrometry The emission spectrographic method of Veillon and Park [52] discussed in Section 13.6.2.3 has been applied to the determination of total sulphur in waste waters. When a solution of a sulphur compound is sprayed into a low-temperature fuel-rich hydrogen-air flame, a blue emission (attributed to molecular S ) is observed near the surface of a cold object placed in the 2 flame. A burner with a borosilicate glass sheath was designed to produce the S emission spectrum with a maximum at 384nm. 2
13.9 Organoboron compounds 13.9.1 Waste waters 13.9.1.1 Inductively coupled plasma atomic emission spectrometry Broekaert and Leis [100] carried out the sequential determination of down to 50µg L-1 boron and several metals (barium, cadmium, copper,
Organoboron compounds
807
iron, manganese, molybdenum, nickel and zinc) by inductively coupled plasma atomic emission spectrometry. This procedure was applied in conjunction with an injection method and the amplified photomultiplier current was used as the analytical signal. 13.9.1.2 Spectrophotometric method Sato [228] examined several hydroxy acids for the extraction– spectrophotometric determination of boron. Mandelic acid was shown to be the most useful. The boron complex obtained is extracted into benzene with malachite green in a simple extraction. Boron is then determined indirectly by measuring the absorption of malachite green in the extract at 633nm. The calibration graph is linear in the range 7.5 × 10-7-1.50 × 105 mol L-1 boron; the apparent molar absorptivity is 6.52 × 104mol-1 cm-1. 13.9.2 Sewage effluents 13.9.2.1 Fluorimetric method An automated fluorimetric method has been applied to the determination [101] of boron in sewage effluents. It is based on the reaction of 4'-chloro-2-hydroxy-4-methoxybenzophenone with boron in 90% sulphuric acid medium to produce fluorescent species. The method permits the determination of different chemical forms of boron, the limit of detection being 1µg L-1. 13.9.3 Mineral waters 13.9.3.1 Ion-exchange chromatography Boron has been determined in mineral wates [102] by ion-exchange chromatography on Dowex 50 WX–8 cation-exchange resin and Dowex 3 weakly basic anion-exchange resin to remove interfering strong electrolytes. Boron is estimated in the column percolate by mass spectrometry. 13.9.3.2 Spectrophotometric and spectrofluorimetric methods Aznarez et al. [227] have described two methods for the determination of boron in water by molecular absorption spectrophotometry with curcumin and by molecular fluorescence with dibenzoylmethane, after extraction of boron into isobutylmethyl ketone with 2-methylpentane– 2,4-diol. The development of the colour or the fluorescence is carried out
808
Determination of elements
in the organic phase used for extraction by addition of curcumin in glacial acetic acid or dibenzoylmethane in isobutylmethyl ketone and phosphoric acid as dehydrating agent. The different conditions for both spectrophotometric and fluorimetric methods have been established. A study has been made of the influence in aqueous solution of several ions as potential interferents. 13.10 Total organic carbon Various techniques have been employed to determine the concentration of dissolved organic carbon (DOC) or total organic carbon (TOC, if unfiltered) in fresh and sea waters. These techniques include wet chemical oxidation [189, 190], photooxidation [191], dry combustion [192], sealed tube combustion [193, 194] and high-temperature combustion [195]. A form of high-temperature catalytic oxidation, which employs a platinum-based catalyst to facilitate oxidation [196], has become the method of choice for the analysis of TOC and DOC in sea water and other natural waters because this method appears to have a greater oxidation efficiency than most of the other methods [197, 198]. Consequently, a large number of high-temperature combustion systems have been built, and several commercial instruments are currently available [199–203]. Even though high-temperature combustion appears to have a greater oxidation efficiency than other techniques, some high-temperature combustion instrumental designs have problems that can give rise to poor reproducibility and poor accuracy. Poor reproducibility and poor accuracy can also arise from contamination during sampling, sample manipulations (e.g. transfers, filtration and acidification), incomplete sparging of inorganic carbon, and sample storage [200, 204–206]. Instrumental problems are mainly related to the sample injection system, memory (or carryover) effects, system blank evaluation, and motion sensitivity. These problems are particularly evident in the analysis of sea water. As a result of the high salt content and the low DOC concentration, ~40µM (0.5ppm) in deep water and 80–90µM (~1ppm) in surface waters, the accurate, precise and artefact-free determination of DOC in sea water has been a major instrumental challenge despite over 30 years of effort [205]. The necessity for accurate and precise determination of DOC in natural waters has increased sharply over the last few years because several large-scale models and major field studies of marine systems have focused on carbon cycling and its relation to global fluxes, climatic fluctuations and food web dynamics. This need has been partly met through recent concerted efforts of the marine community, which involved several workshops and intercomparisons [206–208]. Although precision (and presumably
Total organic carbon
809
accuracy) of DOC analyses of sea water has improved markedly as a result of these efforts, maintaining this high precision is still difficult and labour intensive, especially at sea. Determination of DOC requires two steps. The first step is the conversion of the organically bound carbon to a simple molecular form that can be measured quantitatively. The most common method is the oxidation of organic carbon dioxide by photodecomposition [209, 210], thermal combustion [211], wet oxidation, and pyrolysis methods [212]. As photodecomposition cannot handle refractory non-purgeable organic compounds the wet oxidation method is only applicable for non-volatile carbons, and the pyrolysis method is handicapped by the difficulties in handling large quantities of water involved in the analysis of a low TOC sample; the thermal combustion method is the chosen method as it can oxidize both volatile and non-volatile organic compounds and a large volume of water sample is used during the analysis. The second step of the method is to detect the carbon dioxide evolved during the oxidation of the organic compounds. Carbon dioxide can be directly determined by either non-dispersive infrared absorption spectroscopy [213, 214] or a thermal conductivity detector [215], or indirectly determined after scrubbing in solution by acid–base titration gravimetry [216, 217], ion-exclusion chromatography [218–220] and suppressed ion chromatography [221–223]. Out of the various methods used, non-dispersive infrared absorption spectroscopy methods, conductivity and ion-exclusion chromatography require dedicated equipment, and titrimetry and gravimetry do not have the required sensitivity. Non-suppressed ion chromatography is preferred, it uses normal high-performance liquid chromatography equipment, which is readily available in analytical laboratories, and it has the sensitivity for detecting low level of TOC in given water samples. 13.10.1 Natural waters All natural waters contain inorganic and organic carbon, the ratio depending on the type of water. The relationship between the two is defined as Total carbon (TC) = Total inorganic carbon (TIC) + Total organic carbon (TOC) In almost all natural waters the concentration of inorganic carbon far exceeds that of organic carbon. Depending on the information required, one, or two or all three forms of carbon may be measured. Total inorganic carbon in water consists of the anions HCO - and CO 3
3
810
Determination of elements
together with dissolved carbon dioxide. These fractions are a potential nutrient source and also add to the buffering capacity of the water. Methods for measuring total inorganic carbon normally rely on the use of acid and heat to release the carbon dioxide from the total inorganic carbon, which is quantitatively measured using a number of methods, e.g. infrared analyses, titration and colorimetric determination (Section 13.11). In addition to measuring total organic carbon, instrumental analysers such as the Dohrmann range, which operate by digesting organic carbon in the sample to carbon dioxide with persulphate in the presence of ultraviolet light can also be used to measure total inorganic carbon (Section 13.11). The first stage in the determination of total organic carbon involves the addition of an acid which releases the amount of carbon dioxide equivalent to the inorganic carbon content of the sample, the organic carbon remaining unreacted at this stage. Instruments such as the Dohrmann DC-180 and the DC-85 give a measure of total inorganic carbon as well as total carbon, total organic carbon, purgeable organic carbon and non-purgeable organic carbon. A great variety of methods have been used for the determination of total organic carbon in natural water. Combustion methods include combustion at 850–950°C with or without a catalyst followed by determination of the carbon dioxide produced by non-dispersive infrared analysis [103–110]. Gas chromatography [111] or indirectly coupled plasma atomic emission spectrometry [112] have been used. Combustion followed by conversion of the carbon dioxide produced to methane which is determined gas chromatographically is another technique that has found favour [113–118]. An alternative method of combustion for degrading the sample is to expose it to ultraviolet radiation in the absence [119, 120] or presence [121, 122], of persulphate oxidant, then determine the carbon dioxide produced by non-dispersive infrared spectroscopy. Persulphate or peroxydisulphate oxidation [123] oxidizes organic matter to carbon dioxide which is either determined directly by infrared spectroscopy, or following conversion to methane gas by gas chromatography. Finally, methods have been described based upon ultraviolet spectroscopy of the water sample [125, 126] or by electrochemical means [127, 128]. 13.10.1.1 Thermal combustion methods Several commercially available TOC analysers operate on the principle of ignition of the carbon in the water sample to carbon dioxide which is then determined by non-dispersive infrared spectroscopy. In the Beckman 915 instrument [127, 128], which is now relatively old, the
Total organic carbon
811
sample is initially acidified to release carbon dioxide from any inorganic carbonates in the sample. This carbon dioxide is swept from the sample by aeration. The sample is then injected into the instrument and total organic carbon is determined directly, but any volatile organic carbon is lost. Discrete samples of 10–29µL are injected into a combustion tube at 900°C which contains a small plug of cobalt catalyst. The carbon dioxide produced is measured by a non-dispersive infrared absorptiometer. It is difficult to obtain representative samples, even after homogenization, but satisfactory results can be obtained in the range 10–200mg C L-1 with a precision of about ±3% of the reading if three to six injections are carried out for each sample. In the range 5–10mg C L-1 the percentage error is about ±5% at full scale and ±50% at the lower end of the scale. Below 3mg C L-1 results are very unreliable. The laboratory version is difficult to automate but a version suitable for on-line use is available (the IR915). This instrument permits automatic sampling and analysis of up to 20 samples per hour. Shimadzu [110] supply an instrument marketed in the UK by Smail and Sons Ltd which will determine total carbon and total organic carbon, including the volatile fraction, so that no sample pretreatment, apart from acidification, is required. Continuous sampling is provided by means of a peristaltic pump which allows a more variable and larger sample to be taken. No blocking or poisoning of the catalyst takes place. This is because a regular drip passes into an empty furnace at 850°C, where it drops on to a heated silver block and is broken into tiny droplets which vaporize rapidly. The vapour then passes into the catalyst where oxidation to carbon dioxide is completed. The carbon dioxide is measured by means of an infrared absorptiometer. The sensitivity of the instrument is adjustable in the ranges 0.1, 0.30 and 0.100mg C L-1. Ton and Takahashi [106] have described the Dohrmann DC-90 apparatus which is an improved combustion technique to overcome many of the problems observed with the conventional combustion method. Carrier water was continuously pumped into a ceramic combustion tube provided with a continuous oxygen flow where it was dispersed and instantaneously evaporated. The sample was injected into the carrier water stream. Carbonaceous materials were completely oxidized in the presence of a catalyst and, after condensation of the stream, carbon dioxide in oxygen was measured by a non-dispersive infrared detector. The system blank registered a negligible concentration and the large sample volumes (up to 1000µL) enhanced sensitivity levels as low as 10µg L-1 carbon. In all cases recovery was within 2% of calculated values.
812
Determination of elements
Van Steenderen [107] has discussed the factors which influence the determination of total organic carbon in water. He concludes that samples must be stored in all-glass sealed containers and analysed immediately upon arrival; if this is not possible, the samples must be preserved with silver sulphate solution and stored at 4°C. If dissolved organic carbon is to be determined, the samples must be filtered through a 0.45µm membrane filter previously rinsed with distilled water. Acidification of water samples to remove inorganic carbon is not recommended. The TOC-500 manufactured by Shimadzu [110] operates on the thermal combustion non-dispersive infrared principle. The instrument is fully automated and incorporates a microcomputer. In this instrument the starting and end points of the peak are automatically detected to provide an accurate peak area. Quantitation is made by a two-point calibration curve method. The total organic carbon value is calculated by difference from the total carbon value and the inorganic carbon value. Means, standard deviations and coefficient of variation data are automatically provided. The furnace temperature and baseline stability data are automatically checked and a ready lamp is lit when the instrument is ready for analysis. Automatic baseline irradiation is performed by the instrument. Organic carbon recoveries carried out on a wide range of organic compounds were in the range 98.8–101.8%. Down to 70µg L-1 TOC can be determined with a coefficient of variation of 8.5% (16.5% at the 40µg L-1 level). An alternative to infrared spectroscopy following thermal combustion is gas chromatography [111]. Organic material is oxidized at 900°C in a quartz tube; a three-way electrovalve reverses the flow of the helium carrier gas as the sample is introduced, so that air is not admitted with the sample. The oxidation reaction gases are catalytically converted to carbon dioxide and nitrogen which are measured by gas chromatography with a thermal conductivity detector. A further alternative is inductively coupled plasma emission spectrometry. Rohl and Hoffman [112] discuss problems associated with conventional inductively coupled plasma atomic emission spectrometry methods for the quantification of the dissolved organic carbon content of water using the aspiration of aqueous samples into a nebulizer. These were overcome by converting organic carbon to carbon dioxide by combustion in a heated quartz tube filled with copper oxide. The reaction products were carried with a stream of argon into a torch. The detection limit for carbon was 3ng. A modification of the thermal decomposition infrared approach is to oxidize the organic carbon to carbon dioxide, then reduce this to methane which can be determined gas chromatographically. The
Total organic carbon
813
Tocsin carbon analyser (supplier Phase Separations Ltd, Merseyside, UK) is based on this principle [116–118, 129]. This technique is relatively easy to automate. Total carbon or total organic carbon can be determined, the latter after removal of inorganic carbon by acidification and aeration. Some volatile organic carbons may therefore again be lost. Continuous pumping using a peristaltic pump allows a more variable and slightly larger sample to be taken. The sample passes as a regular drip through a combustion tube packed with cupric oxide at 900°C. The carbon dioxide which is produced then passes through a reduction tube containing a nickel catalyst at 450°C and the methane produced is measured by a flame ionization detector. The combination of a wide-range, linear and sensitive detector and a wide range of sample volumes enables the instrument to determine TOC in the range 0.05–1000mg L-1. There are however, still some operational problems (mainly blocking of the sample inlet to the combustion chamber). The precision obtained with soft-water samples is excellent down to concentrations as low as 0.1mg L-1, a practical limit of detection being about 0.04mg L-1. At 50mg L-1 the error is ±1–2% of reading, at 0.5mg L-1 ±4% of reading and at 0.05mg L1 ± 10% of reading. Stolyarov and Galev [130] described a procedure of direct full hydrogenolysis for determining total and organic carbon in natural water. Carbonate carbon was automatically removed before volatile and non-volatile organic compounds were heated in a helium atmosphere enriched with hydrogen on the surface of a nickel catalyst at 350– 400°C to generate methane. Only minimal modification of the standard serial gas chromatograph design was required to quantify the methane. The reproducibility and accuracy of determination of organic carbon in aqueous solutions were good. 13.10.1.2 Combustion method Hollinger [131] tested the Merz rapid total organic carbon analyser for its performance with lightly contaminated lake water. He found that a significant correlation between total organic carbon and biomass coincided with a blue-green colour. 13.10.1.3 Wet oxidation methods Busse [124] used this technique to determine total organic carbon and compared results obtained with those found by thermal oxidation procedures. Dohrmann have developed their DC.54 ultralow-level analyser which is capable of determining down to 10µg L-1 total organics in
814
Determination of elements
water in 10min. Purgeable organic carbon can also be measured. The principle of the instrument is that 10mL of water sample containing acidified persulphate is transferred to a sparger and then the automatic analysis cycle started. Step one of this cycle consists of a helium purge of the sample transferring carbon dioxide from inorganic sources and purgeable organic carbon to the carbon dioxide scrubber. Here, carbon dioxide is removed with high efficiency and purgeable organic carbon proceeds through the reduction system where it is converted to methane and measured by a flame ionization detector. The detector signal is integrated and the purgeable organic carbon value is displayed directly in the concentration units. In step two, helium pushes the sample through a quartz reaction coil where the persulphate reagent and intense ultraviolet illumination oxidize all remaining organics to carbon dioxide. The sample then passes to a sparger where helium purges and transfers this carbon dioxide to the reduction zone and into the detector. The integrated signal displayed is the total organic carbon. 13.10.1.4 Ultraviolet radiation decomposition procedures A limited amount of work has been carried out on the cold oxidation of organic matter in water by ultraviolet radiation in the absence of chemical oxidants. Oake [119] suggests that photochemical oxidation by ultraviolet radiation may be a practical alternative to hightemperature pyrolysis for the oxidation of organic compounds in the determination of total organic carbon by infrared or flame ionization detection. Lacour and Carroy [120] proposed a cold oxidation method which used ultraviolet radiation to provide the energy for oxidation combined with a new infrared analysis with interferential, narrow-band filters. This apparatus eliminates most of the disadvantages of the hot methods, the temperature never exceeding 80°C This method, it is proposed, overcomes some of the disadvantages of thermal oxidation methods such as encrustation of apparatus, a high degree of corrosion and accumulation of salts in the reactor. The Maihak UV-DOC-UNOR instrument operates on these principles. More commonly used are combinations of ultraviolet degradation with chemical oxidants, such as potassium persulphate and peroxydisulphate. Rosset et al. [122] give a detailed description of the design of automated equipment developed for continuous monitoring of total organic carbon in natural water. The equipment operates by photochemical oxidation of organic matter with peroxodisulphate and ultraviolet radiation, and
Total organic carbon
815
measurement of the carbon dioxide produced using a carbon-dioxidesensitive electrode. Randon et al. [121] describe a method for automatic, continuous total organic carbon measurement. An industrial instrument has been developed which is based on photochemical oxidation of organic matter by peroxydisulphate and UV irradation, followed by measurement of the evolved carbon dioxide using a glass electrode immersed in a solution of sodium bicarbonate and sodium chloride. The carbon dioxide diffuses across a microporous PTFE membrane into the internal electrolyte. The analysis is totally controlled by a microprocessor and yields of 95–100% were obtained for most of the major organic pollutants. Amino compounds were difficult to oxidize and yields for these compounds ranged from 39 to 100%. The Xertex Dohrmann DC-180 low-temperature total organic carbon analyser is a microprocessor-controlled instrument which utilizes ultraviolet-promoted chemical oxidation of carbon to carbon dioxide which is then determined by non-dispersive infrared spectroscopy. Analysis can be completed in 3min with a precision of 5µg L-1. Total carbon, inorganic carbon and purgeable organic carbon can also be measured on the same instrument. The Dohrmann DC-180, in addition, measures non-purgeable organic carbon. Goulden and Anthony [123] have studied the kinetics of uncatalysed peroxydisulphate oxidation of organic material in natural water. By using a large excess of peroxydisulphate and by controlling the pH the oxidation of organic materials can be carried out at atmospheric pressure in a few minutes. 13.10.1.5 Ultraviolet spectrometry Reid et al. [125] have shown that it is possible to determine the concentration of organic carbon in unpolluted stream water by ultraviolet absorption spectrophotometry. However, to avoid serious errors, it is necessary to group data according to flow prior to the calculation of regression equations. Correlations have been found between the ratio of ultraviolet absorptions found at various wavelengths (e.g. 254 and 510nm) with the chemical oxygen demand of the water sample [133, 138]. Ultraviolet spectrometric measurements at various wavelengths (220– 260nm [132], 245–280nm [133] and 200, 260–280, [138] have been used to assess the concentration of total organic compounds in river waters and lake water. Kawai and Musumeci [133] described a method for measuring chemical and biochemical oxygen demand and correlating the results with optical density. After extraction with chloroform and acidification to pH1.0, the ultraviolet absorption is measured at 245 and
816
Determination of elements
280nm. Results of analyses of river water samples from several stations in Brazil are presented. Kato et al. [138] investigated the character of ultraviolet absorption spectra of various river waters. In all districts, there is a high degree of correlation between absorption at 260µm and chemical oxygen demand, BOD, ammonium nitrogen and dissolved oxygen. Absorption at 240nm is correlated with electrical conductivity. From the principal components analysis, applied to the ultraviolet absorption spectra, absorptions at 350nm, 260–280nm and 200nm are useful in classifying water quality. Single and or multiple regression curves are examined to estimate some standard water quality indices. Thompson and Wagstaffe [139] described a method for the rapid detection of organic pollutants in river water utilizing vapour-phase ultraviolet absorption spectrometry. A small amount of a hexane or chloroform extract of the water sample is slowly heated in a graphite tube in a nitrogen atmosphere, while the absorption of the sample is monitored. Peak shape, appearance time, and measurement of other wavelengths enable many substances, including cutting oils, detergents and polychlorinated biphenyls to be characterized. Traces can be completed within 2min. Mrkva [140] has discussed a correlation between chemical oxygen demand and ultraviolet absorption at 254nm, in river water samples. The results show that absorption at 254nm can be used for the routine monitoring of organic pollution. 13.10.1.6 Electrochemical methods Davenport et al. [127] devised a continuous automated electrochemical method. Hauden and Richard [128] described an indirect electrochemical method based on the fact that the potential difference existing at the interface of a metal electrode (ideally platinum), immersed in a solution, corresponds to an electrical double layer acting as a condenser, the capacity of which is reduced when organic matter is adsorbed on the metal surface. 13.10.1.7 Pyrolysis–gas chromatography–mass spectrometry Nakajima [141] attempted the simultaneous determination of total organic carbon and total nitrogen in lake water by pyrolysis–gas chromatography–mass spectrometry. The equipment consisted of a pyrolysis unit, gas chromatography and quadruple mass spectrometer. The relative standard deviation and detection limits were 3.3% for total organic carbon and 5.0% for total nitrogen at 76µg L-1 and 1300µg L-1 respectively. Results obtained agreed well with those obtained separately
Total organic carbon
817
by other methods. The detection limit for nitrogen was too high for application to some lake water samples. 13.10.1.8 Miscellaneous Otson et al. [142] studied the effects of sampling, shipping and storage on total organic carbon levels in water samples containing from 400 to 7500µg L-1 total organic carbon. The sampling, storage and shipping methods used had little effect on total organic carbon levels in tap water and river water samples, and it is concluded that water samples from distant sources can be satisfactorily shipped to a central laboratory for reliable total organic carbon determination. The Department of the Environment [143] have discussed the basic principles and methods of determination using automated instrumental techniques for a range of water quality parameters. The various parameters, starting with biochemical and chemical oxygen demand and including a number of others such as total, dissolved and particular organic carbon etc., are first defined, to demonstrate their relationship and the methodology common to all of them, which is based on hightemperature oxidation followed by measurement of the carbon dioxide produced. Automatic sampling methods are then reviewed, followed by a description of the methods of sample injection, types of oxidation system employed and also the associated detector systems. Procedures are based on non-combustion procedures such as ultraviolet radiation and/or chemical persulphate oxidation combined with analytical finishes based on converting the carbon dioxide produced to methane and estimating this gas chromatographically. The possible interferences and sources of error are considered followed by comments on the performance and reliability of the equipment currently available. A table indicating the manufacturers and types of instrument marketed in the UK with details of the method of operation and probable accuracy are included in this report. Fitzgerald et al. [144] have described an automated system for the determination of total organic carbon. The system uses sequenced, mechanically controlled steps for ampoule manipulation to break open the top of the glass ampoule and to purge each sample of carbon dioxide and nitrogen. A microprocessor data collection system is used to record analytical data. Chi-squared analysis revealed no significant difference between observed and expected values at the 99.9% confidence level for the automated analysis, whereas a parallel manual analysis had a confidence level of only 70%. Peterson and Montgomery [145] have described a method for preparing organic free water for use in total carbon determinations.
818
Determination of elements
De Haan and De Boer [146] have discussed the applicability of light absorption and fluorescence as measures of concentration and molecular size of dissolved organic carbon in humic lakes. Subra et al. [147] discussed the applicability of liquid chromatographic techniques to the determination of total organic carbon in humic acid water extracts. Crane [148] has considered the factors which apply in the selection of a total organic carbon analyser. Qian and Mopper [224] have developed a high-temperature combustion total organic carbon analyser, which has significant design improvements over existing systems. The new injection system directly connects a loop-type autoinjector to the head of the high-temperature combustion column. This connection facilitates the coupling of an autosampler to the injection system. The entire injection process is closed to the atmosphere, thereby improving the precision and eliminating potential contamination during injection. Injections can be made every 3–5min, depending on the injection and inorganic carbon sparging modes used. The high-temperature combustion column was designed without a ‘cold’ zone or dead space at the top. These improvements eliminated the memory (or carry-over ) effect, which is a potential problem in some high-temperature combustion column designs. The high-temperature combustion column is packed with pure quartz beads instead of the relatively expensive platinum-based catalyst, without loss in the oxidation efficiency, as indicated by 100% recovery for various compounds with different refractory properties and by intercomparison with platinum-based high-temperature combustion systems. Precision for sea water is ~±0.6% (r.s.d.) at the 80µM level. Typically, greater than 5000 injections of sea water can be made without significant deterioration of column performance. The effects of column temperature and carrier gas flow rate are reported. Finally, evidence is presented that suggests that there is a relationship between the refractory nature of pure compounds and the peak width. This potential relationship may be a useful tool for quantifying the refractory nature of organic carbon in natural waters. 13.10.2 Potable waters 13.10.2.1 Peroxydisulphate oxidation Van Steenderen et al. [149] give details of the direct automated determination of trace levels of organic carbon in potable water. It involves a wet chemical oxidation technique using silver peroxydisulphate, and measurement of the liberated carbon dioxide by infrared spectrophotometry. The technique is that of continuous flow
Total organic carbon
819
analysis and utilizes an automatic sampler, a multiple-channel proportioning pump and an appropriate analytical system (manifold). Samples are acidified, oxygen segmented and passed through a heated inorganic carbon stripper. The stripped liquid is resampled, mixed with silver peroxydisulphate solution and digested at 70°C. The liberated carbon dioxide is measured by means of infrared spectrophotometry. Complete inorganic carbon removal was not possible. From a 30mg L-1 inorganic carbon solution 0.1% residual remained after stripping. Using a sample volume of 6.3cm3 min-1, full-scale recorder deflection could be obtained for a 1mg L-1 organic carbon standard solution with a relative standard deviation of 0.9%. The lower detection limit of practical significance is 50µg L-1 organic carbon. The analysis rate is 20 samples per hour. Certain synthetic organic materials resisted complete oxidation including the amino acids glycine, 1-alanine, leucine, iso-leucine and valine. 13.10.2.2 Ultraviolet oxidation Ultraviolet oxidation has been used [150] to determine total organic carbon in potable water. The analyser is based on ultraviolet oxidation at low temperature, with an infrared detector for the carbon dioxide evolved. The components are a highly efficient reactor with a submerged ultraviolet tube, a stable, highly sensitive, non-dispersive infrared detector, a microprocessor and a digital display unit for results. The potassium persulphate reagent is fed continuously via a dosing pump and the sample is acidified and oxygenated before injection to the ultraviolet reactor. The calibration and determinations are rapid and simple and the instrument is capable of continuous measurement. Replicate measurements of a sample containing 0.3mg L -1 had a reproducibility of plus or minus 0.011mg L-1. Kehoe [151] discusses the limitations of the total organic carbon method for analysis of water including the limits of detection and the efficiency of conversion of carbonaceous material to carbon dioxide. The correlation of total organic carbon with biochemical oxygen removal and chemical oxygen demand is also considered briefly. 13.10.2.3 Thermal combustion–ion chromatography Fung and Zucheng Wu Dao [225] have discussed a thermal combustion– ion chromatographic method for the determination of total organic carbon in industrial and potable waters. This method utilizes a tube furnace and readily accessable high-performance liquid chromatography equipment. To achieve complete oxidation, persulphate (0.25%) was added to oxidize non-volatile organic compounds in solution and cupric oxide heated at
820
Determination of elements
900°C to convert volatile organic compounds to carbon dioxide, which was scrubbed in a 20mL solution of 50mM potassium hydroxide with 10 drops of butanol added. The carbonate anion obtained was determined by nonsuppressed ion chromatography using 0.6mM potassium hydrogen phthalate as the eluent. Both surfactants and volatile and non-volatile organic compounds commonly found in environmental waters give highly repeatable recoveries close to 100%. The detection limit (S/N = 2) and linear range for a 1L water sample are 2µg C L-1 and 10–2500µg C L-1, respectively, and they can be adjusted using samples ranging from 100mL to 2L. Good repeatability (r.s.d. < 10%) and close to 100% recoveries were obtained for potassium hydrogen phthalate added to real samples such as deionized, mineral, tap, river and sea water. Compared with the ASTM D2579 method, the ion chromatographic method is three orders of magnitude more sensitive, more accurate and more reliable in determining samples with low total organic carbon values and more flexible in adjusting the linear range and sensitivity using variable sample sizes. 13.10.3 Waste waters Soluble and participate organic material can be rapidly converted to the gaseous phase in an appropriate high-temperature reaction environment and the carbon content of the resultant gases can be quantitatively measured and recorded. The commercial instruments available either oxidize carbon in the water sample to carbon dioxide and subsequently measure the carbon dioxide by infrared analysis or alternatively reduce the carbon to methane and measure the methane by a flame ionization technique. No more than 8min per single determination is required for previously standardized instruments. Differentiation between organic and inorganic carbon is relatively easy to obtain with available instrumentation. Based on the organic carbon content of a waste water sample, an approximation of the ultimate oxygen demand of the waste water can be estimated based on stoichiometry. Only very small samples, usually 20–100µL, are required and sampling error may be a problem. Because of the small bore diameter of most total organic carbon (TOC) injection systems, solids must be precluded or reduced in size prior to injection for most instruments. 13.10.3.1 Miscellaneous The determination of total organic carbon in waste water has been reviewed by Busse [152]. Earlier methods [103, 153] for the determination of total organic carbon in waste water involved the use of the Beckman apparatus in which the sample is evaporated at 900–1000°C. The carbon dioxide produced is measured by infrared spectroscopy. These techniques can detect down to 0.5mg L-1 carbon. Dobbs et al. [154] established a correlation
Total organic carbon
821
between the total organic carbon content of waste water and its extinction at 254nm. High correlation coefficients were obtained for samples that contained sufficient carbon to permit accurate determinations. The procedure is not applicable to turbid samples. Becker [155] reviewed methods and equipment available for automated determinations of total organic carbon and inorganic carbon in waste waters. Stover and McCartney [156] characterized industrial waste waters by evaluating biochemical oxygen demand, chemical oxygen demand and total organic carbon and evaluated relationships between these parameters for 30 categories of waste water. Chandler et al. [157] have reported on the feasibility of using measurements of total organic carbon for indirect estimation of biochemical oxygen demand and chemical oxygen demand in waste treatment plants. The results showed a significant correlation between 5day biochemical oxygen demand and total organic carbon. It is concluded that total organic carbon analyses may be used as alternatives to biochemical oxygen demand and chemical oxygen demand tests once the necessary correlations have been determined. Power and Langford [158] have used the thermal lens effect for the thermooptical determination of dissolved organic matter. 13.10.4 Trade effluents 13.10.4.1 Combustion methods Earlier methods for the determination of total organic carbon in trade effluents involved either combustion of the sample in the presence of cupric oxide [159] or digestion with a mixture of concentrated sulphuric and phosphoric acids containing potassium dichromate and potassium iodiate [160]. In all cases complete conversion occurs of organic carbon to carbon dioxide. After drying with magnesium perchlorate the carbon dioxide is determined by a katharometer [159] or by weighing [160]. More recently, automated methods based on exposure of the acidified sample to ultraviolet light have come to the fore, for the determination of total organic carbon in effluents [161]. In the Technicon Industrial Systems Monitor 650 continuous flow analyser, the sample in a silica coil is exposed to ultraviolet light for approximately 2min, which completely breaks down any organic carbon. Any carbon dioxide thus generated is diffused across a gas-permeable membrane to separate it from the reaction matrix, and analysed colorimetrically. The thermal combustion–ion chromatographic method [225] discussed in Section 13.10.2.3 has been applied to the determination of total organic carbon in trade effluent samples.
822
Determination of elements
13.10.4.2 Reduction to methane An alternative method that has been used for the determination of total organic carbon in trade wastes involves injection of the sample into a stream of pure nitrogen which passes over cupric oxide at 850–900°C to convert carbon to carbon dioxide [162]. Hydrogen is then introduced and the mixed gas passes over nickel on firebrick at 350°C to reduce carbon dioxide to methane which is detected by means of a flame ionization detector after chromatography on silica gel. Down to 1mg L-1 carbon can be determined. Croll [163] has investigated this procedure in detail and in conjunction with Phase Separations Ltd (UK) has developed a commercial instrument. The sample is first acidified with nitric acid and then purged with carbon-dioxide-free air in order to remove inorganic carbonates. It is then pumped to the oxidation stage where the sample is vaporized and the organic materials present are oxidized to carbon dioxide. After the addition of nitrogen, the stream is condensed and the nitrogen plus carbon dioxide separated from the water, which is pumped to waste. Hydrogen is added to the gas stream and passage over a nickel catalyst at 450°C reduces the carbon dioxide to methane, which is then measured using a flame ionization detector. 13.10.5 Sewage effluents 13.10.5.1 Combustion methods Various workers have reported on the determination of total organic carbon in sewage [164–166]. In one method [164] a stream of carbon dioxide, alone or mixed with an inert gas is passed through a bed of carbon, at 450–650°C where a small portion of the carbon dioxide is reduced to carbon monoxide and then into a combustion zone containing a catalyst, e.g. platinum, at 500– 1000°C where carbon dioxide reacts with any oxidizable material present in the sample to form carbon monoxide. The increase in carbon monoxide content in the gas stream in the presence of a sample is determined. Tieman and Wagner [165] have described the principles of design and operation of a total organic carbon measuring device with particular emphasis on the need to control the particle size of the suspended solids so as to prevent blockages and false readings. Criteria for the successful performance of a total organic carbon measuring system are outlined, and a system specially developed to meet them and overcome the deficiencies of earlier models is described. The application of this
Total organic carbon
823
Table 13.7 The effect of acidification and membrane filtration on the organic content of settled sewage and humus tank effluent. Results in mg L-1
Source: Reprinted with permission from Van Steenderen [ 107]. Copyright (1976) Erudita Publications Ltd, Johannesburg.
equipment in the operation and process control of activated-sludge plants is described. Van Steenderen [107] found that samples for total organic carbon measurement must at all times be stored in all-glass sealed containers and analysed immediately upon arrival. Failing this, samples must be preserved with silver sulphate solution and stored at 4°C. If dissolved organic carbon is desired, samples must be filtered through a 0.45µm membrane filter previously rinsed with distilled water. The practice of acidifying samples to remove inorganic carbon is not recommended. Van Steenderen [107] analysed sewage and other samples using the Beckmann TOC analyser. This instrument determines organic carbon in water by the difference between total carbon and inorganic carbon. To analyse a sample completely for carbon, two infrared analysers and two automatic sample injectors are necessary, since the instrument is equipped with a dual furnace but only one infrared analyser. He found that over a 2-day period the total organic carbon content of settled sewage samples decreased appreciably, whilst the total inorganic content increased. Acidification of the sample prior to analysis also has a profound effect on reported total oxygen demands (Table 13.7). Van Steenderen [107] used factorial analysis to study the interaction of temperature, preservation and container material simultaneously on the organic carbon content of water samples which had been filtered through a porosity glass filter. Three different temperatures (4, 10 and 30°C) were employed, and preservation was applied at three different concentrations (unpreserved, 1mL saturated AgSO mL-1, 4mL saturated 4 AgSO mL-1). 4 Temperature and container material were the critical parameters affecting the carbon concentration and not preservation. Plastic containers were found to release organic material into solution and this was especially noticeable in the humus tank effluent sample. On the other hand, preservation was the dominant factor when dealing with a chemically unstable and more concentrated organic carbon sample such
824
Determination of elements
as settled sewage where humus tank effluent contained a higher organic carbon content than usual and, as a result, preservation began to play a major role exceeding the effects of temperature and container material. Acidifying of samples had an immediate effect on the organic carbon present and all volatile organic carbon was removed between pH2 and 4. General conclusions are that: samples must be collected and stored in all-glass sealed containers and that preservation of samples must be carried out with silver sulphate solution combined with storage at 4°C. For the determination of dissolved organic carbon, samples must be filtered through a 0.45µm membrane filter previously rinsed with distilled water. Lastly, the acidification of samples in order to remove inorganic carbon from solution is not recommended. 13.11 Total inorganic carbon 13.11.1 Natural waters 13.11.1.1 Spectrophotometric method Automated spectrophotometric methods have been described for the determination of total inorganic carbon in natural waters [167, 168]. Gravelot-Blondin et al. [167] describe a single automated method which uses an adapted Technicon autoanalyser. The sample is segmented with carbon-dioxide-free air, acidified and heated to 37°C, then passed under the dialysis membrane to ensure that a constant fraction of the carbon dioxide diffuses through the membrane into the buffered thymol blue indicator stream. The colour change, measured at 590nm, is proportional to the inorganic carbon content of the water. The method has been successfully used in the range 50–500mg L -1 bicarbonate for total inorganic carbon measurements. Recoveries for 102mg L-1 averaged 98.4% and for 310mg L-1 averaged 101.3%. The coefficient of variation was less than 1%. Linearity of the method is satisfactory in the range 50– 500mg L-1 HCO -. The buffered indicator solution concentrations were 3 selected to obtain optimum linearity and sensitivity. The smaller the volume of buffer used, the higher the sensitivity but then linearity was adversely affected, thus a suitable compromise had to be found. The automated spectrophotometric procedure of Crowther and Moody [168] is appreciably more sensitive than that of GravelotBlondin et al. [167]. Crowther and Moody [168] describe two procedures, capable respectively of determining inorganic carbon in the ranges 0.3–40mg L-1 and 0.01–2mg L-1, whereas the Gravelot-Blondin method [167] is used in the range 50–500mg L-1. The extra sensitivity of the Crowther and Moody [168] method is often needed in the analysis of some types of
Total organic carbon
825
natural water which contain very low concentrations of total inorganic carbon. This method entails conversion of carbon dioxide equilibria products to carbon dioxide. By gas dialysis, a portion of this gas was dissolved in a weakly buffered alkaline phenolphthalein solution, and inorganic carbon was determined by measuring the loss in colour of the indicator. Calibrations were linear within 1% full scale. To estimate the perishability of samples, eight samples were collected simultaneously at one site, and processed at 15min intervals with the first vial analysed 5min after collection. The results were 2.24, 2.24, 2.24, 2.32, 2.24, 2.24, 2.24, 2.28 mg C L-1. Finally, the first vial which had been exposed to the atmosphere for 2h, was reanalysed; the inorganic carbon had decreased from 2.24 to 2.10mg L-1. These results suggest that the procedure is valid for low-level inorganic carbon determinations provided that sample exposure to the atmosphere is restricted. An interference study was conducted on these methods by mixing the test solution with a blank or carbon standard within the manifold. Solution strengths, however, were adjusted such that flow rates and concentrations at the gas dialysers were unchanged. This technique avoided interactions with the atmosphere. The results of the study showed that eight common cations, aluminium, ammonium, calcium, iron (III), magnesium, manganese (II), potassium and sodium, and seven common anions, fluoride, chloride, nitrate, nitrite, phosphate, silicate and sulphate, were not detected as carbons and that carbon spikes were recovered within 2%. At the 50mg L-1 level, humic and tannic acids did not interfere. Sulphide, however, was detected as inorganic carbon, 1mg S2- L-1 was equivalent to 0.6mg C L-1. Although this spectrophotometric system for determining low levels of inorganic carbon required considerable care in preparation of standards and reagents, the resultant data were both precise and relatively free from interference effects. At these levels, however, inorganic carbon is a perishable parameter requiring prompt analysis and limited exposure to the atmosphere. 13.11.1.2 Infrared analysis method Most methods for the determination of total inorganic carbon assume that this source of carbon is present in the sample totally as carbonate and/or bicarbonate. The sample is acidified and the released carbon dioxide measured. Salonen [169] described a simple infrared procedure for the determination of total inorganic carbon, and some gases in water. The sample is injected into a bubbling chamber where gases are stripped from solution and carried to the analyser. Carbonates can be converted to free carbon dioxide by a continuous feed of acid to the bubbling chamber. Excess water is removed with a carrier gas and separated in a condenser.
826
Determination of elements
Since no manipulations are needed, other than to inject samples into the bubbling chamber, the method is very rapid, requiring 20–40s for one sample. The procedure is suitable for routine use and a relative standard deviation is typically about 0.5%. Merkel and Festl [170] determined total inorganic carbon in 30µL samples of natural water with the aid of microcomputers. The analogue output signal of an infrared spectrometer, a linear function of the carbon dioxide concentration, is digitized and transferred on-line to the computer, while the integration step is performed at the end of the run. Equilibrium constants and the pH value are used to calculate the equilibrium distribution of selected inorganic carbon species automatically. 13.11.1.3 Miscellaneous Wilson [171] studied the influence of humic compounds on the titrimetric determination of total inorganic carbon in natural water. He considers the potential effects of humic buffering capacity on total inorganic carbon determinations. The buffer capacity of humic substances is described mathematically and the effects of pH value, carbonate species concentrations and humic concentration on total inorganic carbon are quantified. Humic substances are shown to be poor buffers, even when present in high concentrations. Titrimetric methods for the determination of total inorganic carbon do not, of course, have the sensitivity of the automated spectrophotometric procedure of Crowther and Moody [168]. 13.12 Dissolved organic carbon 13.12.1 Natural waters The measurement of dissolved organic carbon is an important part of many water quality studies. The dissolved organic matter may represent the degradation products of plant or animal life that lives, or has lived, in the water, or alternatively, it may represent pollution by sewage or industrial effluents. Various workers have discussed the determination of dissolved organic carbon in water [172–184]. 13.12.1.1 Potassium persulphate–ultraviolet oxidation Van Steenderen and Lin [172] and Gravelot-Blondin et al. [178] used potassium persulphate combined with ultraviolet irradiation to oxidize
Dissolved organic carbon
827
soluble organic matter in natural water. Gravelot-Blondin et al. [178] automated the procedure. This method uses a modified Technicon autoanalyser. The sample is segemented, acidified, heated to 56°C and sparged with nitrogen. An oxidant mixture of sulphuric acid, potassium persulphate and mercuric nitrate is added before passing the sample through a quartz coil and irradiating it with ultraviolet light. A wetting agent is added and the sample stream is passed through the dialyser into a buffered thymol blue indicator at pH9.6. The colour change is proportional to the dissolved organic carbon concentration. The method is simple and rapid and gives reliable measurements in the 0.5– 29mg L-1 range. The results of recovery tests on this procedure were quite good, a recovery of 99.9–104.8% being obtained for a water sample spiked with potassium hydrogen phthalate. The oxidation efficiency is quite good for a range of organic compounds. 13.12.1.2 Silver peroxydisulphate oxidation Ehrenberger [179] determined traces of dissolved organic carbon in natural water after wet oxidation using a silver-ion-activated solution of peroxydisulphate in sulphuric acid. The decomposition product is combusted on silver permanganate. The carbon dioxide formed during conversion is adsorbed on a molecular sieve, then desorbed by heat and measured by thermal conductivity. The output of the detector is digitally encoded for read-out and further processing. 13.12.1.3 Ultraviolet oxidation Baker et al. [180] have described a procedure for the measurement of dissolved organic carbon and particulate organic carbon in river water and chalk streams. The method is based on the oxidation of organic carbon to carbon dioxide by ultraviolet radiation, the carbon dioxide produced being measured by a non-dispersive infrared gas analyser. The latter was also used in the semiautomatic measurement of particulate organic carbon by a combustion method. Surveys of the Piddle and Frome river systems indicated dissolved organic carbon concentrations of less than 2.0mg L-1 as typical of unpolluted chalk streams, but in streams draining acid heathland significantly higher concentrations were found. The rapid increase of dissolved organic carbon concentration during river spates is discussed. Particulate organic carbon concentrations of 0.03– 0.04mg L-1 were found in chalk spring waters. The relation of the measurement of dissolved and particulate organic carbon flux to estimates of the energy budgets of stream ecosystems is discussed and published methods for the automatic measurement of dissolved organic carbon are reviewed.
828
Determination of elements
An automatic ultraviolet oxidation method has been described [181] for the determination of down to 10µg L-1 of dissolved organic carbon in water from the Great Lakes. The inorganic carbonate is removed in a heated packed column, the organic carbon is oxidized by one of two methods (ultraviolet irradiation or silver-catalysed peroxydisulphate at 95°C) and the resulting carbon dioxide is measured by an infrared analyser. The silver-catalysed peroxydisulphate method was shown to be the more convenient and precise, but it does not completely oxidize all materials in water; in a large number of natural water samples analysed the level of organic carbon oxidation averaged 97%. The limit of detection for carbon in 10µg L-1 and the analysis rate is 20 samples per hour. 13.12.1.4 Ultraviolet spectroscopy De Haan et al. [182] showed that there was a direct correlation between absorption at 250nm and the concentration of dissolved organic matter in a humic lake water. 13.12.1.5 Miscellaneous Nakajima [173] simultaneously determined dissolved organic carbon and phosphorus in natural waters using flame ionization and photometric detection. Cation interference is eliminated by passage of the sample through a cation-exchange resin, and an ultrasonic nebulizer is used to introduce the sample to the equipment. The long-term relative standard deviation was 5.8% for dissolved organic carbon and 5.2% for dissolved phosphorus. Detection limits were 90µg L-1 and 30µg L-1 respectively. Leenheer [175] gives details of a system for the preparative isolation and fractionation of dissolved organic carbon in natural waters by adsorption on a series of resins. The various organic fractions can then be characterized by infrared spectra. Sontheimer and Kuhn [183] outlined the parameters employed for a broad indication of the nature and extent of organic pollution of surface water systems. As bulk parameters, either dissolved organic carbon or chemical oxygen demand could be used to provide an indication of the total level of organic matter. Group parameters included the determination of the content of heteroatoms such as chlorine, bromine, sulphur and nitrogen together with the measurement of biochemical oxygen demand as an indicator of biodegradability. Wilhelms [184] investigated the possibility of carrying out simultaneous determinations of dissolved organic carbon and inorganic carbon compounds in river and lake waters. He used a thin-film dissolved organic carbon analyser for total inorganic carbon
Dissolved organic carbon
829
determination (carbonate, bicarbonate and dissolved carbon dioxide). A stream of oxygen was passed through the solution and the resulting mixture of oxygen and carbon dioxide led to an infrared analyser. The dissolved organic carbon determination was performed by the use of irradiation of a thin film of the sample with ultraviolet light from a mercury discharge lamp, thus oxidizing the organic carbon to carbon dioxide which was also determined by infrared absorption. Details involved are described for the calibration of the system and the automation of the measuring equipment with the aid of a double-beam infrared spectrometer and a double-pen recorder. Hara et al. [185] have described a flow method for the determination of organic carbon which utilizes a washing system and is equipped with a potentiometric carbon dioxide sensitive gas electrode. To avoid interference from nitrogen dioxide, sulphide and hydrogen sulphide originating from organic or inorganic nitrogen and sulphur compounds in the water sample it is recommended that the sample is adjusted to pH5 before analysis. 13.12.2 Potable waters 13.12.2.1 Miscellaneous Servais et al. [186] give details of two procedures for the determination of biodegradable dissolved organic carbon in potable water. In both procedures the sterile filtered sample is reinoculated with a natural bacterial population from the same region as the sample and incubated in the dark for 10 days at 20°C. In one procedure, dissolved organic carbon is measured until a plateau is reached, the difference between the initial and final dissolved organic carbon being taken as the biodegradable organic carbon. In the other procedure, bacterial biomass and mortality rate are followed and integrated flux of mortality during the incubation period calculated and divided by the growth yield to give an estimate of biodegradable dissolved organic carbon. 13.12.3 Trade effluents 13.12.3.1 Persulphate oxidation Oxidation with potassium persulphate is commonly used to determine dissolved organic carbon effluents [187]. Oxidation is carried out in the presence of silver ions for 30min at 50–80°C. The carbon dioxide formed is carried by an inert gas over heated cupric oxide and absorbed in potassium hydroxide solution. Inorganic carbon dioxide is determined
830
Determination of elements
first by acidification of the sample and collection of the carbon dioxide. In both instances after precipitation of the carbonate with barium chloride the unconsumed potassium hydroxide is titrated. 13.13 Particulate organic carbon 13.13.1 Natural waters 13.13.1.1 Combustion methods Hulsmann and Hengst [188] used wet oxidation by potassium persulphate in sealed ampoules to estimate particulate organic carbon. Inorganic carbon was first removed from the sample prior to reaction with persulphate. Krambeck et al. [134] measured small quantities of particulate carbon in lake waters by an automated furnace combustion infrared procedure. The whole sequence of operations was controlled with the aid of an AIM65 desktop computer; the system was successfully operated for routine analysis of samples of lake water with particulate organic carbon values of 100–300µg L-1 carbon; a single analysis took 8min. The relative standard deviation was about 1%. Fresh deionized water contains undetectable amounts of dissolved inorganic carbon. The absence of significant reagent or other blanks makes it possible to perform determinations down to 1µM concentration of dissolved inorganic carbon. Prior to the combustion of organic gases the carrier gas, with gases evolved from the sample, was conducted through a soda lime tube to remove carbon dioxide. A test was made to assess whether the absorber allows quantitative passage of the reduced organic gases present in lake waters. The sum of separate estimates of gaseous organic carbon, and dissolved inorganic carbon in lake waters agreed well with that obtained by combusting all evolved gases (without prior absorption). In no individual case were the two results significantly different. Thus the absorption procedure does not seem to cause losses of reduced organic gases. In 10 bicarbonate standard samples, with dissolved inorganic carbon levels distributed evenly between 0.8 and 19mg L-1, the mean coefficient of variation was 0.47% (SD = 0.16). For the same samples, the standard deviation of the calibration coefficients was only 0.7% of the mean, thus indicating very good linearity. The precision of the method was so slightly dependent on the injected volume and of the flow rate of the carrier gas, that it is unimportant for most practical purposes.
References
831
Hilton et al. [135] have described an automated method, based on the Erba Science 1106 elemental analyser for the determination of particulate carbon and nitrogen in natural waters. 13.13.1.2 Miscellaneous Salonen [136] compared different glass-fibre and silver-metal filters for the determination of particulate organic carbon. He puts forward a theory that the varying characteristics of different filters, or pore sizes, appreciably modify the results of particulate organic carbon determinations, making comparison of published concentrations unreliable. Salonen compares silver-metal filters and different types of glass-fibre filters, and seeks to find a filter which would have biologically meaningful cut-off size of particles. The most retentive glass-fibre filters were able to retain almost all bacteria from the water of an oligotrophic lake; they prove to be quite near to the ideal. Silver filters provide similar retention, but because of their high blank values, price, and lower filtration speed and capacity, they are not able to compete with glassfibre filters in practical work. 13.13.2 Rain water 13.13.2.1 Digestion method Ogren et al. [137] have described a method for the determination of elemental carbon in rain water in which organic and elemental carbon are separated by prefiltration, digestion and volatilization. Elemental carbon masses as low as 10µg are determined with ±50% accuracy. References 1 Wegman, R.C.C. and Greve, P.A. Science of the Total Environment 7 235 (1977) 2 Fritschi, E., Frishi, G. and Kussmani, H. Z. Wasser Abwasser Forsch. 11 165 (1978) 3 Brandt, G. and Kettrup, A. Int. J. Environ. Anal. Chem. 31 129 (1987) 4 Stachel, B., Lahl, U., Schroer, W. and Zeshmar, B. Chemosphere 13 303 (1984) 5 Genther, J., Lunde, G. and Steinnes, E. Anal. Chim. Acta 108 137 (1979) 6 Ahnoff, M. and Josefsson, B. Analytical Chemistry 46 658 (1974) 7 Hasanen, E. and Manninen, P. Chemosphere 16 969 (1987) 8 Kuhn, W., Fuchs, F. and Southheimer, H. Z. Wasser Abwasser Forsch. 10 192 (1977) 9 Van Steenderen, R.A. Labor Pract. 29 380 (1980) 10 Jager, W. and Hagenmaier, H. Z. Wasser Abwasser Forsch. 13 66 (1980) 11 McCahill, M.P., Conroy, L.E. and Maier, W. Journal of Environmental Science and Technology 14 201 (1980) 12 Leddicote, G.W. and Navarette, M. Revte. Soc. Quim. Mex. 12 223A (1968)
832
13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49
Determination of elements
Sadisivan, S. and Anand, S.J.S. Ind. J. Environ. Health 21 10 (1979) Knapp, G. and Spitz, H. Talanta 16 1631 (1969) Jeronimo, G. Revta. Assoc. Bioquim. Argent. 38 207 (1973) Pesavento, M. and Profumo, A. Analyst (London) 110 181 (1985) Erdey, A. and Kozmutza, K. Acta Chim. Hung. 69 9 (1971) Williams, D.T., Coburn, A. and Bonesi, J. Journal of the Environment International 10 39 (1984) Nulton, C.P., Haile, C.L. and Redford, D.P. Analytical Chemistry 56 598 (1984) Kaj, L. and Solyom, P. Vatten 44 121 (1988) Trifonova, N.A. Trudy Isvest. Bio. Unutr. Vod. Akad, Nauk, S.S.S.R. 18 247 (1968) Ref. Zh. Khim . 19GD (12) Abstr. No. 129 248 (1969) Smart, M.M., Reid, F.A. and Jones, J.R. Water Research 15 919 (1981) Lennox, L.J. and Flanagan, N.J. Water Research 16 1112 (1982) Harwood, J.E. and Huyser, D.J. Water Research 4 539 (1970) Sybandim J.C. and Hafkamp, W. H O 8 333 (1975) Lowry, J.H. and Mancy, K.H. Water2 Research 12 471 (1978) Solorzano, L. and Sharp, J.H. Limnology and Oceanography 25 754 (1980) Picat, B., Hammou, N., Casellas, C, Jourdan, M.F. and Bontoux, J. Water Supp. 6 1 (1988) Mancy, B.A., Miller, M.C. and Welzel, R.G. Limnology and Oceanography 16 71 (1971) Wood, O. Analytical Abstracts 15 7628 (1968) Nydahl, F. Water Research 12 1123 (1978) Mertens, J., Van der Winkel, P. and Massert, P. Analytical Chemistry 47 522 (1975) Johansen, H.S. and Middelboe, V. International Journal of Applied Radiation and Isotopes 27 591 (1976) Leicknam, J.P., Figdor, H.C., Keroe, E.A. and Muehl, A. Int. J. Appl. Radiat. Isotopes 19 235 (1968) Karlsson, L. and Middelboe, V. Isotopes and Radiation in Soil–Plant Relationships Including Forestry, p. 211, IAEA, Vienna (1972) Fiedler, R. and Proksch, G. Anal. Chim. Acta 78 1 (1975) Middelboe, V. International Journal of Applied Radiation and Isotopes 29 752 (1978) Fiedler, R. and Proksch, G. Anal. Chim. Acta 78 29 (1975) Nelson, D.W. and Sommers, L.E. Journal of Environmental Quality 4 465 (1975) Sjostrom, L., Radestrom, R., Carlberg, G.E. and Kringstad, A. Chemosphere 14 1107 (1985) Jenkins, D. Prog. Water Technol. 8 31 (1977) Adamski, J.H. Analytical Chemistry 48 1194 (1976) O’Connor, B. and Miloski, P. Unpublished work, Suffolk County Department of Environmental Control, Suffolk, UK, October (1974) Armstrong, F.A.J., Williams, P.M. and Strictland, J.D.H. Nature (London) 211 481 (1966) Stevens, R.J. Water Research 10 171 (1975) Tenny, A.M. Autom. Anal. Chem. 580 (1966) Moore, R.T. and McNulty, J.A. Environmental Science and Technology 3 741 (1969) Albat, M., Picat, B., Philpot, J.M. and Bentoux, J. Rev. Franc. Sci. L’eau 1 359 (1982) Elkei, O. Anal. Chim. Acta 86 63 (1976)
References
833
51 Irie, T., Sakino, H., Hamada, S. and Komine, K. Journal of Japan Sewage Works Association 22 66 (1985) 52 Veillon, C. and Park, J.Y. Anal Chim. Acta 60 293 (1972) 53 Jirka, A.M., Carter, M.J., May, D. and Fuller, F.D. Environ. Sci. Technol. 10 1038 (1976) 54 Bowman, G.T. and Delfino, J.J. J. Water Pollut. Contr. Fedr. 54 1324 (1982) 55 Jenkins, D. Progress in Water Technology 8 31 (1977) 56 Starck, B. and Haapala, K. Valten 41 54 (1985) 57 Daughton, C.G., Jones, B.M. and Sakaji, R.H. Analytical Chemistry 57 2326 (1985) 58 Croll, B.T., Tomlinson, T. and Whitfield, C.R.W. Analyst (London) 110 861 (1985) 59 Haga, K. and Harado, Y. Journal of Japan Sewage Works Association 25 74 (1988) 60 Helfgott, T. and Mazurek, J.S. Progress in Water Technology 8 433 (1977) 61 Cabrera, F., Diaz, E., Toca, C.G. and De Arambarrui, P. Water Research 16 1061 (1982) 62 Pescas, J.T., Faugere, J.G. and Vanbesier, M. Tribune du Cebedeau 34 189 (1981) 63 Environment Canada Analytical Methods Manual, Inland Water Directorate Water Quality Branch, Ottawa, Ontario, Canada (1974) 64 Jeffries, D.S., Kieken, F.P. and Jones, D.E. Water Research 13 275 (1979) 65 Goossen, J.T.H. and Kloosterboer, J. Analytical Chemistry 50 707 (1978) 66 Pekukhova, L.A. Biol. Vnutr. Vod. Inform. Byull. (16) 71 (1972) Ref. Zhur Khim 19GD (22) Abstract No. 22G110 (1972) 67 Zaiyon, L. and Limin, W. Talanta 33 98 (1986) 68 Harwood, J.E., Van Steerderen, R.A. and Kuhn, A.L.A. Water Research 3 425 (1969) 69 Grasshoff, K.Z. Analytical Chemistry 220 89 (1966) 70 Osburn, Q.W., Lemmel, D.E. and Downey, R.L. Environ. Sci. Technol. 8 363 (1974) 71 Harvey, H.W. J. Mar. Biol. Ass. 27 337 (1948) 72 Lee, G.F., Clesceri, N.L. and Fitzgerald, G.P. Air Water Pollut. 9 715 (1965) 73 Menzel, D.W. and Corwin, N. Limnol. Oceanog. 10 280 (1965) 74 Goulden, P.D. and Brookbank, P. Anal. Chim. Acta 80 183 (1975) 75 King, D.E. and Fellin, P. Ontario Ministry of the Environment, Laboratory Services Branch, Data Quality Report Series, Section 1. The Water Quality Laboratories, Data Quality Summary (1975) 76 Lennox, L.J. Water Research 13 1329 (1979) 77 Ichinose, N., Kania, H., Nakamura, K., Shimizu, C., Kurobira, H., Okamato, K. and Inui, T. Anal. Chim. Acta 15 345 (1984) 78 Hashimoto, S., Fujiwara, K. and Fuwa, K. Limnol. Oceanogr. 32 729 (1987) 79 Frimmel, F.H. and Grenz, R. Fresenius Z. Analyt. Chemie 327 11 (1987) 80 Ikeda, N., Yoshikawa, M., Murakami, S. and Kunicka, J. Journal Radioanalytical and Nuclear Chemistry 122 27 (1988) 81 Olsen, O., Gotterman, H.L. and Clymo, R.S. (editors) Chemical Environment in the Aquatic Habitat. Proceedings of an IBP Symposium, Amsterdam 1– 16October, 1966 . NV Noord-Haollansche, Uitgevers Moat Schappi, Amsterdam (1967) 82 Ishizuka, T., Nakajima, K. and Sunahra, H. Anal. Chim. Acta 121 197 (1980)
834
Determination of elements
83 American Public Health Association, American Water Works Association and Water Pollution Control Federation. Standard Methods for the Examination of Water and Waste Water, APHA, Washington DC, 14th edn, p. 466(1975) 84 Aoyagi, M., Yumamasa, Y. and Nishida, A. Anal. Chim. Acta 214 229 (1988) 85 Crowther, J., Weight, B. and Weight, W. Anal. Chim. Acta 119 313 (1980) 86 Shirai, T. Water Research 17 1721 (1983) 87 Ebina, J., Tsutsui, T. and Shirai, T. Water Research 17 1721 (1983) 88 Hasomi, M. and Sudo, R. Int. J. Environ. Stud. 27 267 (1986) 89 Henricksen, A. Analyst (London) 95 601 (1970) 90 Hasomi, M. and Sudo, R. J. Waste Water. 21 1433 (1979) 91 Muhlhauser, H.A., Soto, L. and Zahradnik, P. Int. J. Environ. Anal. Chem. 28 157 (1979) 92 Nriagu, J.O. and Hem, J.D. In Sulfur in the Environment, J.O. Nriagu (ed.) Wiley Interscience, New York, Part 2, pp. 211–270(1978) 93 Shriner, D.S. and Henderson, G.S. J. Environ. Qual. 7 392 (1978) 94 Mitchell, M.J., Landers, D.H. and Brodowski, D.F. Water Air Soil Pollut. 16 351 (1981) 95 David, M.B., Mitchell, M.J. and Nakas, J.P. Soil Sci. Am. J. 46 847 (1982) 96 Landers, D.H., David, M.B. and Mitchell, M.J. Int. J. Environ. Anal. Chem. 14 245 (1983) 97 Chakraborti, D. and Adams, F. Anal. Chim. Acta 109 307 (1979) 98 Boulengue, J., Ciabrini, J.P., Fouillac, G.C., Michard, G. and Ouzounian, G. Chem. Geol. 25 19 (1979) 99 Brull, E.E. and Goulden, G.S. Anal. Chim. Acta 110 167 (1979) 100 Broekaert, J.A.C. and Leis, F. Anal. Chim. Acta 109 73 (1979) 101 Afghan, B.K., Goulden, P.D. and Ryan, J.F. Water Research 6 1475 (1972) 102 Gorene, B., Marcel, J. and Tramsek, G. Mikrochimica Acta 1 24 (1970) 103 Axt, G. Vom Wasser 36 328 (1969) 104 Jones, R.M. and Dageford, A.F. ISA Trans. 7 267 (1968) 105 Helms, J.W. US Dept. of the Interior Geological Survey Report 4373 . Rapid measurement of organic pollution by total organic carbon and comparisons with other techniques (1973) 106 Ton, N. and Takahashi, Y. Am. Labor. 17 48 (1985) 107 Van Steenderen, R.A. Water South Africa 2 156 (1976) 108 Van Steenderen, R.A. Water South Africa 2 126 (1976) 109 Madier, D. Vom Wasser 36 246 (1969) 110 Shimadzu Corporation International Marketing Division, Shinjuku, Building 1–1 Nishi-Shinjuku 2-chrome, Shinjuku-ku, Tokyo 163, Japan 111 Bertoni, R. and Maestroni, G.J.M. Memorie de l’Instituto Italiano di Idrobiologia Dott. Marco de Marchi, 34 131 (1977) 112 Rohl, R. and Hoffman, H.J. Fresenius Z. Analyt. Chemie. 322 439 (1985) 113 Takahashi, V., Moore, R.T. and Joyce, R.J. Int. Labor. Sept./Oct. 21 (1972) 114 Takahashi, V, Moore, R.T. and Joyce, R.J. Am. Labor. 4 31 (1972) 115 Croll, B.T. Chem. Ind. 9 386 (1972) 116 Cropper, F.R., Heinekey, D.M. and Westwell, A. Analyst (London) 92 436 (1967) 117 Cross, B.T. Chemy Ind. 9 386 (1972) 118 Cross, B.T. Measurement of total organic carbon (TOC) in waters and effluents, Technical Memorandum TM 86, Water Research Centre, Medmenham, Marlow, Bucks (1973)
References
835
119 Oake, R.J. A review of photo-oxidation for the determination of total organic carbon in water, Technical Report TR 160 Water Research Centre Environmental Protection: Water Research Centre, Medmenham, UK (1981) 120 Lacour, G. and Carroy, A. Eau, Industrie, Nuisances, No. 75 59 (1983) 121 Randon, G., Dutang, M., Sassiat, P., Caude, M. and Rosset, R. Rev. Franc. Sci. L’eau 3 53 (1984) 122 Rosset, R., Caude, M., Sassiat, P. and Dutang, M. Int. J. Environ. Anal. Chem. 13 19 (1982) 123 Goulden, P.D. and Anthony, D.H.J. Analytical Chemistry 50 953 (1978) 124 Busse, H.J. Z. Wasser Abwasser Forsch. 9 82 (1976) 125 Reid, J.M., Cresser, M.S. and MacLeod, D.A. Water Research 14 525 (1980) 126 Lowden, G.F. J. Water Pollut. Contr. Fed. 50 10 (1976) 127 Davenport, R.J., Wynveen, R.A. and Cooper, W.J. J. Am. Water Works Ass. 73 555 (1981) 128 Hauden, D. and Richard, Y. Progr. Water Technol. 7 41 (1975) 129 Croll, B.T. The determination of total organic carbon in water, Technical Memorandum TM 124, Water Research Centre, Medmenham, UK, May (1976) 130 Stolyarov, B.V. and Galev, E.E. Soviet J. Water Chem. Technol. 10 57 (1988) 131 Hollinger, W. Z. Wasser Abwasser 12 193 (1979) 132 Sheppard, C.R.C. Water Research 11 979 (1977) 133 Kawai, H. and Musumeci, M.R. Revista 39 81 (1979) 134 Krambeck, H.J., Lampert, W. and Bredic, H. Fachz. Laborat. 25 2009 (1981) 135 Hilton, J., Lishman, J.P., Mackness, S. and Heaney, S.I. Hydrobiologia 141 269 (1986) 136 Salonen, K. Hydrobiologia 67 29 (1979) 137 Ogren, J.A., Charlson, R.J. and Grobilicki, P.J. Analytical Chemistry 55 1569 (1983) 138 Kato, Y., Shirakata, Y. and Ziganic, M. Japan J. Water. Pollut. Res. 2 45 (1979) 139 Thompson, K.C. and Wagstaffe, K. Analyst (London) 104 668 (1979) 140 Mrkva, M. Water Research 17 231 (1983) 141 Nakajima, K. Water Research 20 233 (1986) 142 Otson, R., Williams, D.T., Bothwell, P.D., McCullough, R.S. and Tate, R.A. Bull. Environ. Contam. Toxicol. 23 311 (1979) 143 Department of the Environment/National Water Council Standing Committee of Analysts, HM Stationery Office, London. Methods for Examination of Waters and Associated Materials (1980) 24 pp. The instrumental determination of total organic carbon, total oxygen demand and related determinands (1979) 144 Fitzgerald, L., Montgomery, J.R. and Holt, J.K. J. Autom. Chem. 6 80 (1984) 145 Peterson, G.N. and Montgomery, J.R. Anal. Chim. Acta 113 395 (1980) 146 De Haan, H. and De Boer, T. Water Research 21 731 (1987) 147 Subra, P., Foucault, A. and Dorset, R. Int. J. Environ. Anal. Chem. 31 197 (1987) 148 Crane, G.A. Am. Labor. 20 51 (1988) 149 Van Steenderen, R.A., Basson, W.D. and Van Duuren, F.A. Water Research 13 539 (1979) 150 Landre, J. Eau, Industrie, Nuisances No. 71 25 (1985) 151 Kehoe, T.J. Environ. Sci. Technol. 11 137 (1977) 152 Busse, H.J. Z. Wasser Abwasser 9 17 (1976) 153 Bauer, L. and Shmitz, W. Vom Wasser 36 383 (1970) 154 Dobbs, R.A., Wise, R.H. and Dean, R.B. Water Research 6 1173 (1972)
836
Determination of elements
155 Becker, W.J. Gas-u.-Forschung (Wasser und Abwasser) 120 217 (1979) 156 Stover, E.L. and McCartney, D.E. BOD results that are believable. In Fourth Annual Meeting of Pollution Control Association of Ponca City, Oklahoma (1984) 157 Chandler, R.L., Shaugnessy, J.C. and Blane, F.C. J. Water Pollut. Contr. Fed. 48 2791 (1976) 158 Power, J.F. and Langford, C.H. Analytical Chemistry 60 842 (1988) 159 Ehrenberger, F. Z. Analytical Chemistry 267 17 (1973) 160 Stevenson, M.E., Cabrera, B.E. and D’ Itro, F.M. Environ. Set. Technol. 5 799 (1971) 161 DeLiddo, J., Conelta, A. and Dorsheimer, W. Int. Labor. 60, October (1984) 162 Cropper, F.R. and Heinekey, D.M. Analyst (London) 94 484 (1969) 163 Croll, B.T. The measurement of total organic carbon in waters and effluents, Technical Memorandum TM 86, Water Research Association, Medmenham, UK, July (1973) 164 Stenger, V.A. and Clayton, E. British Patent, 1,169,373 19/12/66 165 Tieman, A. and Wagner, B. Korrespond. Abwasser 27 658 (1980) 166 Nelson, K.H., Lysyj, I. and Nagano, J. Water Sewage Works 117 14 (1970) 167 Gravelot-Blondin, L.R., Van Vleit, H.R. and Schoones, B.W.H. Water South Africa 6 135 (1980) 168 Crowther, J. and Moody, W.B. Anal. Chim. Acta 120 305 (1980) 169 Salonen, K. Water Research 15 403 (1981) 170 Merkel, B. and Festl, U. Z. Wasser Abwasser Forsch. 17 153 (1984) 171 Wilson, D.E. Archiv. Hydrobiol 87 379 (1979) 172 Van Steenderen, R.A. and Lin, J.S. Analytical Chemistry 53 2157 (1981) 173 Nakajima, K. Water Research 18 555 (1984) 174 Gloor, R., Leidner, H., Wuhrmann, K. and Fleischman, T. Water Research 15 457 (1981) 175 Leenheer, J.A. Environ. Sci. Technol. 15 578 (1981) 176 Smart, P.L., Finlayson, B.L., Rylands, W.D. and Ball, C.M. Water Research 10 805 (1976) 177 Mitchell, D.G., Aldous, K.M. and Canelli, E. Analytical Chemistry 49 1235 (1977) 178 Gravelot-Blondin, L.R., Van Vleit, H.R. and Mynhardt, P.A. Water South Africa 6 138 (1980) 179 Ehrenberger, F. Z. Wasser Abwasser Forsch. 8 75 (1975) 180 Baker, C.D., Bartlett, P.D., Farr, I.S. and Williams, G.I. Freshwater Biol. 4 467 (1974) 181 Goulden, P.D. and Brooksbank, P. Analytical Chemistry 47 1943 (1975) 182 De Haan, H., De Boer, T., Kramer, H.A. and Voerman, S. Water Research 16 1047 (1982) 183 Sontheimer, H. and Kuhn, W. Fresenius Z. Analyt. Chemie. 324 225 (1986) 184 Wilhelms, A. Z. Wasser Abwasser Forsch. 15 53 (1982) 185 Hara, H., Okabe, Y. and Kitagawa, T. Analytical Chemistry 64 2393 (1992) 186 Servais, P., Ballen, G. and Hascoet, M.C. Water Research 21 445 (1987) 187 Schmid, H.G. IT Fachz Lab. 15 1105 (1971) 188 Hulsmann, A. and Hengst, P. Hydrobiol. Bull. 14 135 (1980) 189 Duursma, E.K. Neth. J. Sea Res. 1 1 (1961) 190 Menzel, D.W. and Vaccaro, R.F. Linmol. Oceanogr. 9 138 (1964) 191 Armstrong, F.A.J., Williams, P.M. and Strickland, J.D.H. Nature (London) 221 481 (1996) 192 Skopintsev, B.A. Mar. Hydrophys. Inst. 19 1 (1960) 193 Fry, B., Saupe, S., Hullar, M. and Peterson, B.J. Mar. Chem. 41 187 (1993)
References
194 195 196 197 198 199 200 201 202 203 204 205 206 207 208 209 210 211 212 213 214 215 216 217 218 219 220 221 222 223 224 225 226 227 228
837
Alperin, M.J. and Martens, C.S. Mar. Chem. 41 135 (1993) Sharp, J.H. Mar. Chem. 1 211 (1973) Sugimura, Y. and Suzuki, Y. Mar. Chem. 24 105 (1988) Sharp, J.H., Suzuki, Y. and Munday, W.L. Mar. Chem. 41 253 (1993) Koprivnjak, J.F., Blanchette, J.G., Bourbonniere, R.A. and Clair, T.A. Water Research 29(1) 91 (1995) Tanoue, E. Earth, Planet, Sci. Lett. 111 201 (1992) Peltzer, E.T. and Brewer, P.G. Mar. Chem. 41 243 (1993) Martin, W.R. and McCorkle, D.C. Limnol. Oceanogr. 38 1464 (1993) Chen, W. and Wangersky, P.J. Mar. Chem. 41 167 (1993) Cauwet, G. Mar. Chem. 47 55 (1994) Sharp, J.H., Peltzer, E.T., Alperin, M.J., Cauwet, G., Farrington, J.W., Fry, B., Karl, D.M., Martin, J.H., Spitzy, A., Tugrul, S. and Carlson, C.A. Mar. Chem. 41 37 (1993) Wangersky, P.J. Mar. Chem. 41 61 (1993) Tupas, L.M., Popp, B.N. and Karl, D.M. Mar. Chem. 45 207 (1994) Hedges, J.L. and Farrington, J. Mar. Chem. 41 5 (1993) Sharp, J.H., Benner, R., Bennet, L., Carlson, C.A., Fitzwater, S.E., Peltzer, E.T. and Tupas, L. Mar. Chem. 48 91 (1995) Edwards, R.T., McKelvie, I.D., Ferett, P.C., Hart, B.T., Bapat, J.B. and Koshy, K. Anal. Chim. Acta 26 287 (1992) Van Steenderen, R.A. and Liu, J.S. Analytical Chemistry 53 2157 (1981) Montgomery, H.A.C. and Thom, N.S. Analyst (London) 87 689 (1962) Test Methods for Total and Organic Carbon in Water, American Soc. Testing Materials, Philadelphia, PA, 1985; ASTM D2579-85 Huber, S.A. and Frimmel, F.H. Analytical Chemistry 63 2122 (1991) Zhao, Z. Huaxue Shijie 32 215 (1991) Kieselbach, R. Analytical Chemistry 26 1312 (1954) Seligson, D. and Seligson, H. Analytical Chemistry 23 1877 (1951) Maxon, W.D. and Johnson, M.J. Analytical Chemistry 24 1514 (1952) Tanaka, K. and Fritz, J.S. Analytical Chemistry 59 708 (1987) Krelling, J.R. and DeZwann, J. Analytical Chemistry 58 3028 (1986) Technical Note, No. 7. The Wescan Ion Analyzer, Wescan Instruments, Santa Clara, CA, (1984) Application Brief No. 2004 , Waters Chromatography Division of Corporation, Milford, MA (1986) Okada, T. and Kuwamoto, T. Analytical Chemistry 57 829 (1985) Cochrane, R.A. and Hillman, D.E. J. Chromatogr. 241 392 (1982) Qian, J. and Mopper, K. Analytical Chemistry 68 3090 (1996) Fung, Y.S. and Zucheng Wu Dao, K.L. Analytical Chemistry 68 2186 (1996) Cheung, Y.Y., Kirkbright, G.F. and Snook, R.D. Anal. Chim. Acta 140 213 (1982) Aznarez, J., Bonilla, A. and Vidal, J.C. Analyst (London) 108 368 (1983) Sato, S. Anal. Chim. Acta 151 465 (1983)
Chapter 14
Oxygen demand parameters
14.1 Chemical oxygen demand Apart from the standard potassium chromate digestion procedure, several other procedures have been used for the determination of the chemical oxygen demand of natural waters including continuous flow injection analysis [1], combustion–infrared spectroscopy [2], titration with nitrate [3, 4], spectrophotometry [5] and potentiometric titration [6]. In the direct potentiometric method for determining chemical oxygen demand, Dugin et al. [4] used a galvanic cell with no liquid junction. Reversibility of the system was achieved by adding cerium (IV) to the working solution. A titanium silicate glass electrode allowed measurement of cell potential E and temperature in strong acid solution h and at temperatures over 100°C. A number of potentiometric results for various compounds were determined and the method allowed kinetic information to be obtained as well as chemical oxygen demand values. The error of the procedure did not exceed 3% and the lower limit of detection was 10mg L-1. 14.1.1 Natural waters
14.1.1.1 Titration methods Baughman et al. [7] refluxed the sample with excess acid dichromate in the presence of silver sulphate and titrated unconsumed dichromate potentiometrically (tungsten reference electrode and platinum indicator electrode) with 0.025N ferric solution using automatic titration. Kelkenberg [8] modified the standard acid dichromate procedure by using 88% sulphuric acid and heating the sample for 2h at 150°C in an open vessel. In this way he was able to obtain near quantitative recoveries for acetic acid, ethyl alcohol and aromatic compounds.
Chemical oxygen demand
839
Various workers [9–11] have discussed the elimination of the interference effect by chloride in the determination of chemical oxygen demand. Wagner and Ruck [10, 11] discuss three methods for overcoming interference: (1) quantitative oxidation of chloride ions to chlorine – with certain precautions – and correction of the result of the analysis for the calculated amount of oxidizing agent needed for this reaction; (2) use of mercuric sulphate as sequestering agent for chloride ions (the limits of this method are discussed); (3) elimination of the chloride ions from the sample prior to the oxidation step in the form of hydrogen chloride by diffusion in a closed chamber to avoid losses of volatile oxidizable constituents. In this latter method [11] chloride is removed in the form of hydrogen chloride from a strongly acidic solution of the sample. It was shown that in doing so – even with higher chloride concentrations – no losses of acetic acid occur. From solutions of acetic acid test substance the observed interference by nitrate ions can be traced to the fact that the reduction of nitrates to nitrite-nitrogen by chloride ions is enhanced in the presence of higher sulphuric acid concentrations and the nitrites also contribute to the chemical oxygen demand value. As a preventative measure in the case of water samples containing nitrate ions the hydrogen chloride diffusion can be carried out using weaker sulphuric acid solutions and, if necessary, a longer duration of the analysis can be accepted. Messenger [12] compared determinations of chemical oxygen demand in open flasks and sealed borosilicate glass ampoules and discuss the advantages and disadvantages of the sealed tube technique. Various workers have discussed the elimination of mercury salts [13– 15] and the replacement of the silver oxidation catalyst by other catalysts. No satisfactory replacement for the silver catalyst has yet been found, although it may be possible to use reduced amounts [16]. No catalyst has been found to have universal oxidation properties [17]. Walter and Adams [18] have presented an updated version of the standard method for chemical oxygen demand in which chloride interference is suppressed by the use of a silver catalyst instead of mercury. The workers described how to eliminate interference by sulphur dioxide in the determination of chemical oxygen demand. Ogneau [19] has discussed improvements to the standard dichromate method for the determination of chemical oxygen demand. Le Clair et al. [20] have discussed the effects of chloride on the determination of chemical oxygen demand which involved the addition of large excesses of sulphate to the sample prior to analysis. Automatic titrimetry has been applied to the determination of chemical oxygen demand [21].
840
Oxygen demand parameters
14.1.1.2 Coulometric method Fleet [22, 23] described a procedure based on stoichiometric reactions between dichromate ions and hydrogen peroxide added in excess in acid medium to produce oxygen which is measured coulometrically using a three-electrode oxygen analyser [23]. Silver, mercuric ions, chloride, phosphate and urea do not interfere. Nitrite is removed by sulphamic acid and chloride is masked with mercuric ions. 14.1.1.3 Spectrophotometric method Some work has been carried out on the automation of the chemical oxygen demand method [24–28] using autoanalysers. In the autoanalyser technique [28] the sample is treated with concentrated sulphuric acid containing potassium dichromate in the presence of silver sulphate and mercuric sulphate, in a coil 24m long immersed in a bath at 65°C. Determinations on samples containing up to 500mg L-1 chemical oxygen demand, confirmed the validity of the method. Corrections are necessary in the presence of appreciable amounts of chloride ion. Up to 80 determinations per day could be performed. Thomas and Muningda [29] review recent modifications to the standard method for chemical oxygen demand determination and report their own investigations of the operating conditions. They show how it is possible to decrease the quantities of reagents, in particular the amount of silver sulphate catalysts, by up to 10 times, and shorten the oxidation time, by increasing the ratio of the sulphuric–phosphoric acid mixture to the other reagents. They developed a rapid micromethod which involves the use of 2mL samples, 3mL of concentrated sulphuric acid, containing 6.6g L -1 silver sulphate, and 1mL 0.25N potassium dichromate. The reaction mixture is heated at 160°C for up to 2h in the presence of mercuric sulphate and then determined spectrophotometrically at 600nm. 14.1.2 Estuarine and sea waters The chemical method for the determination of the chemical oxygen demand of non-saline waters involves oxidation of the organic matter with an excess of standard acidic potassium dichromate in the presence of silver sulphate catalyst followed by estimation of unused dichromate by titration with ferrous ammonium sulphate. Unfortunately, in this method, the high concentrations of sodium chloride present in sea water react with potassium dichromate producing chlorine:
Chemical oxygen demand
841
6Cl- + Cr2O72+ + 14H → 2Cr3+ + 3Cl2 + 7H2O Consequently the consumption of dichromate is many times higher that that due to organic material in the sample. To complicate matters any amines in the sample consume and release chlorine in a cyclic process leading to high chemical oxygen demands. RNH → NH + CO + H O 2 2 2 2 NH + + 3Cl → NCl + 3Cl- + 4H4 2 3 2NCl → N + 3Cl 3
2
2
Also, the addition of silver sulphate causes precipitation of silver chloride which in the presence of organic compounds, is neither completely nor reproducibly oxidized. This method, whilst being applicable to estuarine waters of relatively low chloride content, would present difficulties when applied to highly saline estuarine and sea waters of low organic content. 14.1.2.1 Miscellaneous Southway and Bark [30] discussed the problem of chloride interference in the determination of the chemical oxygen demand (COD) of estuarine waters. They investigated three processes for removing chloride from the test solution, namely removal by chemical precipitation and filtration, removal as chlorine in an oxidation stage, and removal by a ligand exchange process. This last process was found to produce the most satisfactory results. Takimoto et al. [31] conducted a survey on the monthly variations of COD and total organic carbon (TOC) in coastal marine waters of the northern part of Hiroshima Bay. The COD/TOC ratio varied from 0.23 to 2.2; the smaller value was observed when chlorophyll a concentration was high and vice versa. This variation was found to be due to the imperfect degradation of carbonaceous compounds in the COD method determinations. COD values for samples with high concentration of particulate organic materials tended to give smaller COD/TOC ratio. 14.1.3 Waste waters The COD test is a relatively simple, straightforward wet chemistry method for determining the oxygen equivalent of waste water constituents that are susceptible to oxidation by a strong oxidizing agent. The procedure for determining the oxygen demand of a waste water sample is relatively rapid, requiring about 3h and has the advantage of being able to digest colloidal and participate material. Unfortunately, the
842
Oxygen demand parameters
COD test will reflect the oxygen equivalent of organic compounds that are not readily biodegradable and does not completely oxidize benzene, toluene, pyridine and similar compounds. However, the COD test is commonly used for monitoring the performance of industrial waste water treatment systems. COD data cannot be expected to correlate well with biochemical oxygen demand (BOD) data primarily because the two tests are measuring different aspects of the oxidizable constituents of waste water. In addition, the limitations of the BOD test further hamper meaningful correlations. The COD test has been semiautomated for handling large numbers of samples and several modifications have been proposed to shorten the time required to minimize chemical requirements. 14.1.3.1 Titration methods The generally used method [32–40] for estimating the chemical oxygen demand of waste waters involves heating the sample with an excess of acidic potassium dichromate in the presence of silver ions to catalyse the reaction and with mercuric ions to prevent or reduce interference from chloride ions. Excess dichromate is then determined by titration with standard ferrous ammonium sulphate to a suitable Redox indicator end point. The standard UK procedure [35] employs both titrimetric and a spectrophotometric (at 440nm) finish. The determination of chemical oxygen demand in waste waters by a sealed-tube procedure, using silver nitrate to suppress chloride interference, has been described by Ballinger et al. [32]. The procedure employed the standard reflux conditions commonly used in water authority laboratories. The sealed-tube chemical oxygen demand procedure [36] offers several advantages over the reflux procedure, notably a saving in bench space and cost of equipment. In spite of these advantages and general comparability with reflux procedures [32, 35] the sealed-tube procedure has not been adopted as a standard method. The Department of the Environment’s Standing Committee of Analysts (UK) decided not to endorse a sealed-tube chemical oxygen demand procedure because of safety considerations. Thermal degradation of the tube caps and liners is also found to occur. This creates difficulties in unscrewing caps and occasionally causes contamination of samples, particularly if an oven is used to heat the tubes. The apparatus described by Lloyd [37] overcomes the problems cited above and offers several advantages over the silver nitrate reflux procedure. It is not necessary to use a weaker solution for chemical oxygen demand levels of 200mg L -1 or less. The suppression of chloride interference is improved and lower concentrations of silver nitrate are used.
Chemical oxygen demand
843
In this procedure [37] samples are digested at 150°C in a glassstoppered flask using springs to retain the stopper. The use of sealed flask conditions offers two advantages over reflux conditions; simplified apparatus and experimental procedure and improved suppression of chloride interference. This procedure is similar to the standard procedure in accuracy and reproducibility of results. Within-batch relative standard deviation in waste water analysis ranged from 2.2% at 60mg L-1 of chemical oxygen demand to 0.8% at 380mg L-1 of chemical oxygen demand (four degrees of freedom). Total relative standard deviation in analysis of 300mg L-1 chemical oxygen demand potassium hydrogen phthalate solution was 1.0% (eight degrees of freedom). These results did not differ significantly (p = 0.05) from data obtained using the standard procedure or the silver nitrate reflux procedure. Ballinger et al. [32] have also studied mercury-free methods for the determination of the chemical oxygen demand of waste waters. Akesson and Lind [34] evaluated two simplified methods for the determination of the chemical oxygen demand in waste waters. These methods both used capsules prepared with chemicals in advance and then fed with 2mL samples. The methods were compared with the standard method. Both the simplified methods had advantages with respect to time, space, safety and ease of handling, but had disadvantages in regard to costs, precision and accuracy. For waste waters with a high chemical oxygen demand (above 100mg L-1) the capsule methods afforded values about 10% higher than the standard method, while in the low chemical oxygen demand range (under 100mg L-1) the values were lower. 14.1.3.2 Spectrophotometric methods Automation of the chemical oxygen demand test has been discussed [33]. The waste water sample is mixed in the autoanalyser with an oxidant solution comprising potassium dichromate and 75% sulphuric acid. After heating to 145°C the mixture is treated with hydrogen peroxide to produce molecular oxygen, which is measured, after separation of the aqueous phase, and absorption of carbon dioxide by means of a sensor based on a porous catalytic silver electrode. Nitrite and chloride interfere in the procedure. The oxygen demand index of [41] a sample is determined by heating it for 20min at 100°C with potassium dichromate in the presence of a strong acid to oxidize the organic matter. Silver sulphate is used as a catalyst and mercuric sulphate is added to prevent interference from chloride solution. The extinction of the resulting solution is measured at 600nm. The test was found to be useful for
844
Oxygen demand parameters
providing a rapid estimate of the biochemical oxygen demand of waste water provided that enough data were collected at each sampling point. Correlation of results between the oxygen demand index and the 5-day dilution biochemical oxygen demand was inferior to that for the biochemical oxygen demand and the standard chemical oxygen demand but was superior to that for the biochemical oxygen demand and total organic carbon. 14.1.3.3 Flow injection analysis Korenga and Ikatsu [40] have described a flow injection analysis technique with an acidic dichromate carrier, for the continuous determination of chemical oxygen demand in waste water samples. The system compares well with other methods and has the advantages of being cheap and simple. Appleton et al. [42] also described a flow injection system in which a reagent stream containing potassium dichromate and sulphuric acid was merged with a distilled water carrier into which the sample was injected. The combined stream then flowed through a suitable reactor where the sample was oxidized and hexavalent chromium was reduced to the trivalent state; the absorption of chromate was measured downstream at 445nm. Results obtained on several types of waste water by this method and by the standard method are compared. Interference by chloride could be prevented by adding mercuric sulphate. 14.1.4 Trade effluents Work on the determination of the chemical oxygen demand of trade effluents is summarized in Table 14.1. 14.1.5 Sewage effluents 14.1.5.1 Titration methods Numerous workers have studied various aspects of the measurement of the chemical oxygen demand of sewage samples [7, 18, 63–68] (Table 14.2). The use of conventional reflux methods versus sealedtube digestion versus micro and semimicro open-tube methods in the acid dichromate method for determining chemical oxygen demand of sewage has been the subject of debate [69, 70]. Best and De Casseres [70] evaluated the micro sealed-tube method using synthetic standards and a variety of sewages. Comparison of the results with the conventional reflux method indicates that the sealed-
Table 14.1 Determination of chemical oxygen demand of trade effluents
Table 14.1 Continued
Chemical oxygen demand
847
Table 14.2 Acid dichromate methods for measurement of chemical oxygen demand of sewage
tube method using a reagent consisting of potassium dichromate (10– 126g), concentrated sulphuric acid (167mL) and mercuric sulphate (33.3g) is reliable and accurate and suitable for the analysis of large numbers of samples. It is, however, subject to the changes inherent in any sealed-tube method. It was observed that the sealed-tube method oxidized organic compounds more efficiently than the reflux method. Thus ethanol was digested to the extent of 94% in sealed tubes and only 54% in the reflux method. A dilution of at least 1:5 was necessary to obtain satisfactory precision at either 590nm or at 440nm, which is ? for the chromium (VI) max species. The absorption at 440nm decreased with increasing COD, but the sensitivity was about five times greater at 440nm than at 590nm, especially after the necessary 1:5 dilution of the digested sample. In these circumstances similar results were obtained by the manual titration and the spectrophotometric methods. Limits of detection were similar for both methods (13mg L-1 open tube versus 10mg L-1 sealed tube). It is concluded that the open-tube method has all the advantages of the sealed-tube method such as savings in space and cost of chemicals and increased sample throughput. In addition it has the advantage that possible operational hazards are greatly reduced. The precision data and recovery tests
848
Oxygen demand parameters
indicate that the method is as precise and accurate as the sealed-tube method even for volatile compounds. Goodwin et al. [72] report on studies of the kinetics of the dichromate oxidation of dissolved organic matter of the type that occurs in crude sewage and sewage effluents. A system of characterization of oxidizable material was derived comprising a set of four separate parameters calculated from second-order reaction kinetics. Application of this process to chemical oxygen demand determinations allows solutions with the same chemical oxygen demand but different composition to be differentiated, owing to the changes in parameter values associated with changes in the relative proportions of different reacting species. An automated method for the measurement of chemical oxygen demand which has been applied to the analysis of sewages is the Westhoff Hydromat COD manufactured by Bochum, Germany [73]. In this instrument sample and reagents are fed continuously into the instrument by means of a peristaltic pump. The sample is mixed in the ratio 1:1 with concentrated sulphuric acid and fed to a series of separators where chloride is removed as hydrochloric acid by a vigorous air stream from the membrane pump. The stripped liquid is mixed with potassium dichromate and silver sulphate in concentrated sulphuric acid, segmented with air and passed through a glass coil in the oxidation reactor at 160°C. The digested fluid is then mixed with ferrous– ferric solution and the change in potential and hence the relative COD measured in a Redox cell. Using this method a standard deviation of 7.5mg L-1 was obtained for a sample of settled sewage with a chemical oxygen demand of 626mg L-1 (i.e. r.s.d. 1.2%). 14.2 Total oxygen demand The total oxygen demand (TOD) technique consists of determining the quantitative amounts of oxygen required to convert the oxidizable constituents in a waste water to end products under near ideal combustion conditions. Again, instrument response time is 2–3min and detection limits are on the order of 5mg L-1 for oxygen-demanding materials. Acceptable particulate size is dependent on the instrument purchased and most waste waters will require physical disintegration prior to injection. A TOD determination may include the oxygen demand of inorganic materials such as sulphides and ferrous ions. Both TOC and TOD instrumentation can be provided with on-line, semicontinuous determinations of soluble waste water strength. Total oxygen demand is a parameter which measures the oxygen demand of a water sample by combustion at 900°C of oxidizable matter,
Total oxygen demand
849
at a defined oxygen level, during the combustion process. For the user, there exist three important requirements: 1 2 3
The possibility of measuring total oxygen demand over a wide range of values. The times per measurement should be minimal. Results should be attainable without recourse to a calibration curve, i.e. they should be direct readings.
These requirements impose strict conditions with regard to the design of equipment for measuring total oxygen demand. Since complete combustion of the oxidizable matter must be assured, within a short period of time, the sample volume itself must also be small – typically, 10µL (larger volumes tend to generate excessive volumes of water vapour and thus undesirable pressure pulses in the actual measuring system). Hence the quantity of oxygen consumed and measured will only be a few micrograms. Furthermore, to avoid possible uncertainties of measuring small differences between two large quantities, the oxygen content of the gas prior to combustion must be low; and, for good reproducibility, it must also be constant. Finally, since large excesses of oxygen can give different levels of oxidation when substances containing nitrogen and sulphur are oxidized, the oxygen concentration should be matched to the probable oxygen demand of any practical sample, and the value can vary by as much as 100. These requirements are met, for example, by the Philips total oxygen demand meter. This instrument utilizes two well-known electrochemical relationships: Faraday’s law, which relates to the electrochemical transport of elements, and Nernst’s law, which is concerned with electrochemical potentials at boundaries. It is known that, at elevated temperatures and under the influence of an electrical current, oxygen can be transported through specially prepared zirconium oxide and that, in the electrolyte state and in the presence of oxygen, zirconium oxide will generate an electrical potential at its surface. Furthermore, when there exists a difference in the oxygen concentration, there will be a measurable potential difference. In the Philips total oxygen demand meter the zirconium oxide is specially prepared and is in the form of two series-connected tubes or cells, through which a nitrogen carrier gas is allowed to flow. Each cell is provided with two pairs of annular (internal and external) electrodes and the whole assembly is maintained at about 600°C. One pair of electrodes conducts an electrical current through the zirconium oxide wall, thereby transporting oxygen from the external atmosphere
850
Oxygen demand parameters
through the cell wall, into the nitrogen carrier gas. The other pair is used to measure the potential difference over the zirconium oxide. Several nitrogen-containing substances were tested by Voorn and Marlow [74] using the Philips total oxygen demand meter. Their results strongly suggest that hydrogen and carbon were oxidized and nitrogen was not. Some oxidation of combined nitrogen to nitric oxide may take place when measurements are made low in a range, so that an excess of oxygen is available for combustion. Although poorer correlation was obtained with organic substances containing nitrogen, where chemical oxygen demand values are lower than total oxygen demand values (with chemical oxygen demand the combined nitrogen is converted to ammonia and many nitrogencontaining substances are not oxidized by the dichromate to the theoretical degree), it is firmly believed that the total oxygen demand value measured by the Philips instrument closely approaches the true oxygen demand due to natural degradation, because nitrogen is the end product of the natural nitrogen cycle. Sulphur mainly oxidizes to sulphur dioxide. 14.2.1 Natural waters 14.2.1.1 Combustion methods Almost without exception, commercially available TOD meters are based on the principle of combustion in air of a known weight of sample at 850– 900°C during which carbon and nitrogen in the sample are oxidized. This is followed by measurement of the oxygen depletion of the oxygen flowing through the combustion furnace by means of a zirconium oxide fuel cell (Table 14.3); e.g. C H OH + 1/2O → 3CO + 4H O 3 7 2 2 2 C H O N + eO → aCO + 1/2bH O + 1/2dN a
b
c
d
2
2
2
2
The TOD is then calculated from the measured oxygen depletion. An exception is the apparatus described by Von Becker [75] in which the samples are combusted at 1100°C with air instead of oxygen as the carrier gas, oxygen partial pressure measurements being made with a zirconia electrode. Difficulties encountered are discussed and the need for a more systematic investigation of the effect of process variables on the results is stressed. Removal of any excess carbon dioxide prior to the determination of the residual gas partial pressures is shown to be essential for reproducible results. Various workers have discussed the Philips TOD meter [76–78].
a
Notes Precision Scientific Co., Chicago, USA; dt. Vertregung: Classen + Co., 2000 Hamburg. b,c Ionics, Inc., Watertown, Mass., USA; dt. Vertregung: Schwing Verfahrenstechnik KG, 4133 Neukirchen-Vluyn. d Philips Elektronic Industrie GmbH, 3500 Kassel. e Astro Ecology Corp., Houston, Texas, USA; dt, Vertregung: Kontron Technik GmbH, 8057 Eching.
Table 14.3 Summary of commercial TOD meters
852
Oxygen demand parameters
An alternative instrument, based on similar principles, is the Ionics 225 or Ionics 1326 analyser [79, 80] which employs a platinum–lead fuel cell instead of zirconium oxide. The performance characteristics of the latter instrument are such that precision is much better in the middle and upper working ranges (0– 400mg L-1 and 0–1000mg L-1 TOD) than in the lowest range (0–50mg L-1 or 100mg L-1). It has been demonstrated that the instrument will accept samples that contain high levels of dissolved solids and also suspended solids, provided that in the latter case the samples are pretreated so as to reduce the size of the solids to less than 200µm. Laffont et al. [81] and Capdeville and Rogues [82] have studied changes in the forms of carbon, nitrogen, sulphur and principal mineral elements, during measurement of total oxygen demand. Bicarbonate and carbonates are reduced and cause negative interference, while cations are oxidized and cause a positive error in the results. Organic and ammoniacal nitrogen are partially oxidized by high-temperature catalytic action in the total oxygen demand meter. Nitrous and nitric nitrogen cause negative interferences, which must be corrected for. The simultaneous determination of total oxygen demand and total nitrogen demand has been achieved by interfacing the equipment with a nitric oxide analyser. Using a chemiluminescent nitric oxide detector, precisions have been achieved of 1% r.s.d. at the 1mg L-1 total organic nitrogen level [83]. 14.2.1.2 Miscellaneous Fukunaga et al. [84] in addition to measurements of biochemical oxygen demand and chemical oxygen demand, also assessed organic pollution levels in river waters using measurements of TOD and total organic carbon. The ratio of total TOD to soluble TOD was found to be about 1.5. There was a close correlation between total oxygen demand and biochemical oxygen demand, and chemical oxygen demand and suspended solids in rivers. The concept of elemental oxygen demand has been discussed by Nakajima et al. [85]. This is defined as the oxygen demand of each element in the pollutants. They discuss the relation between elemental oxygen demand and total oxygen demand. An equation was developed to represent this relationship and was shown to give good agreement with observed total oxygen demand values for various inorganic and organic compounds. Total oxygen demand analysers have been shown to give more reliable analysis of hydrocarbons in natural water than chemical oxygen demand measurements [86].
Total oxygen demand
853
There have been several reviews of methods for the determination of oxygen demand parameters, including total oxygen demand [87–89]. 14.2.2 Waste waters 14.2.2.1 Miscellaneous Voorn and Marlow [74] applied the Philips total oxygen demand meter technique to waste water samples. They found an excellent correlation with chemical oxygen demand for a range of pure substances containing carbon, hydrogen and oxygen. Marty and Aim [90] have described an automatic determination of the total oxygen demand of waste water. The method gives rapid results and correlates well with chemical oxygen demand and biochemical oxygen demand. Measurements of corrections are made for nitrogen compounds which are the principal interferents. Other workers who have discussed the determination of total oxygen demand include Wells [91] and Ravenscroft [92]. 14.2.3 Trade effluents Voorn and Frampton [93] have given a description for the Philips TOD meter and have discussed its application to industrial effluents. They confirm its usefulness for continuous monitoring of effluents. Arin [87] discusses total oxygen demand meters and their effectiveness for monitoring industrial effluents, although he emphasizes that their limitations must be appreciated before they can be applied successfully. 14.2.4 Sewage effluents 14.2.4.1 Titration method Stones [94] discusses the use of silver-catalysed dichromate to determine the strength of sewage and to assess the performance of treatment plants. By using an appropriate modification it is possible to eliminate interference by chlorides and thus measure the total carbonaceous oxygen demand of sewage. The relation between this and the biochemical oxygen demand and total oxygen demand is discussed. 14.2.4.2 Electrochemical method Cummings [95] has described an automated procedure for determining total oxygen demand in which two cells are set up separated by an anion-
854
Oxygen demand parameters
exchange resin. Each cell contains a nickel electrode. The sample is put in the cathode cell and adjusted to pH10 in the anode cell. Upon passage of a current an amount of oxygen is liberated the partial pressure of which is related to the total oxygen demand of the sample. 14.2.4.3 Miscellaneous Musselwhite [96] has given results of tests on the performance of a commercially available total oxygen demand meter and on factors affecting performance. The tests showed that the figure given by the meter is a measure of the oxygen equivalent of oxidizable matter minus the oxygen liberated under the conditions of the test. It is concluded that this would be useful for sewages and non-nitrified and denitrified effluents but useless for nitrified effluents and waters containing appreciable amounts of oxidized nitrogen. 14.3 Biochemical oxygen demand The definitive UK method for determining 5-day biochemical oxygen demand has been published [97]. Biochemical oxygen demand tests are historically and routinely used for a great many purposes, including some for which the test is not well suited. In the majority of cases, the test is performed by the dilution (bottle) method and the incubation time is 5 days. The results are often designated BOD . For example, BOD tests are used at waste water 5 5 treatment plants to assess influent and effluent strength. Loading and design criteria for biological treatment processes are frequently based on kilograms of BOD per day. Even the quality of the receiving water is 5 often evaluated on this basis, although in this application there is seldom, if ever, a rational justification for the 5-day incubation in terms of real-world conditions. The most serious and obvious limitation of BOD as a parameter for 5 process control or effluent monitoring is its long response time. Good reproducibility in this test is attained only when the sample tested contains a variety of readily biodegradable compounds, mostly of natural origin, or only slightly modified from their natural sources. When the sample contains substantial proportions of natural or synthetic products that require energy input before they become readily degradable, precision suffers seriously. A simple example is acetate. One molecule of adenosine triphosphate is required per molecule of acetate to enter the Krebs cycle. Samples high in acetates usually show a lack of reproducibility in the standard BOD test because of a variable lag 5 period before the onset of rapid oxidation. On the other hand, a sample that contains a preponderance of readily degradable components tends
Biochemical oxygen demand
855
to exhibit a more repeatable course of oxygen utilization and hence a more reproducible value after an arbitrarily fixed time. There are many cases for which a 5-day BOD value alone has little meaning. A different incubation period, a series of points at various incubation periods (that is, a ‘BOD curve’), use of an adapted microbial culture, a technique other than the bottle dilution method or a combination of such modifications is often far more informative. For many kinds of samples, a BOD curve may be needed for intelligent interpretation of a 5-day value. 14.3.1 Natural waters Hewitt et al. [98] used a re-aeration technique to establish oxygen uptake curves for samples of water from four streams in New Jersey, and a stepwise non-linear regression approach was used to calculate rate constants and ultimate BOD values for equations in which the order of the reaction was assumed to be 1, 1.5, 2, 2.5, 3, 3.5 and 4. Although the error mean square was reduced with increase in reaction order from 1 to 4, this was accompanied by approximately a two-fold increase in ultimate BOD values. It is concluded that the ultimate BOD values calculated at the higher reaction orders are probably unrealistically high. Klein and Gibbs [99] have described a graphical method for calculated BOD. This involves plotting a graph of dissolved oxygen remaining after incubation in a series of dilutions, versus volume of sample added; the data are then converted to BOD by means of an equation. This procedure has several advantages. No significant difference has been found in 5-day and 20-day BODs measured on natural waters in 60mL and 300mL incubation bottles. The volume to surface area ratio of the bottle did not affect the BOD. Raff [100] carried out experiments on the effect of different quantities of allylthiourea employed as a nitrification inhibitor during the standard BOD determination. No interference due to allylthiourea on the carbaceous BOD was detectable and the differences between the inhibited and non-inhibited values for BOD of samples of river water from the Regnitz river could be correlated with the ammonia concentration in the samples, and confirmed published values of the oxygen demand of ammonia in aqueous solutions. Rigin et al. [101] determined the BOD of natural waters by a method which involves incubation of the samples at a constant partial pressure of oxygen equal to its atmospheric partial pressure. The oxygen is obtained by electrolysis of a 0.5M solution of potassium sulphate, using a platinum grid anode. The amount of oxygen consumed is determined by an automatic coulometric recorder. The error associated with the measurement of the mass of oxygen absorbed is less than
856
Oxygen demand parameters
0.02% and the BOD results are less scattered than those of other methods. This is attributed to the constant conditions for oxygen absorption. Several oxidation vessels may be attached to a single electrolysis unit, which allows simultaneous determinations in parallel. 14.3.1.1 Automation of BOD procedure Several attempts have been made to partially [102–104] or fully automate [102–106] the BOD procedure. Crowther et al. [102] give details on semiautomated equipment involving probes and dissolved-oxygen meters connected to a teletype machine producing dissolved-oxygen data in computer compatible form. The large-scale incubation facility is also described. Kohne et al. [106] evaluated an automated method for rapid BOD determination employing the dilution principle, designated the BOD-M3 method in several laboratories. It was claimed to be suitable for measurements within the BOD range of 2–5000mg L -1. The method involved introducing the sample at a controlled rate into a continuous flow reactor which operated on a special fluidized bed principle; the rate of oxygen uptake was determined by oxygen electrodes and the strength of the sample calculated from the stepwise increase in the oxygen uptake curve. Results of comparative tests between the standard BOD dilution 5 method and the rapid continuous method are reported, with analysis of interlaboratory variation and reproducibility of the respective test methods. The reproducibility of the BOD-M3 method was between 95 and 98% and exceeded that of the standard BOD test procedure. 5 Gantner [105] describes an apparatus for automatic recording of the 5-day oxygen demand of water samples, in which the oxygen is produced electrolytically from a cell connected to a constant voltage supply. The current to the electrolysis cell is controlled by a mercury relay which responds to changes in the pressure of oxygen in the flask. By recording the time for which the current is flowing and integrating the time–current curve a measure of the oxygen consumption is obtained. 14.3.1.2 Electrometric method Strigel [107] has discussed an electrometric process for the determination of BOD and its change time. These measurements are performed in a closed flask using an oxygen electrode.
Biochemical oxygen demand
857
14.3.1.3 Effects of experimental parameters, (temperature, partial pressure of oxygen, and humidity) Steinecke [108–110] has carried out detailed investigations of several aspects of the BOD method. He discusses available methods for the direct measurement of BOD values with the aid of a supply of gaseous oxygen, either at constant or diminishing partial pressure, and the accuracy of the results obtained. Laboratory apparatus of different types is described with the aid of diagrams and the effects of operating conditions, such as temperature, volume of sample, partial pressure and humidity, on the different apparatus are considered. Details of the operating ranges and the relative accuracies obtained under standard conditions are discussed. Steinecke [109] outlined several procedures for testing the performance. Leakproof connections are essential, and the manner in which the system can be tested to ensure that this condition is fulfilled is indicated for differing types of equipment. Methods are discussed for the calibration of selected system components including the burette, manometer and current regulating device. In further work, Steinecke [110] discusses methods of calibrating the overall response, involving the measurement of the oxygen uptake by volumetric and barometric methods. Ranchet et al. [111, 112] investigated the effect of two storage methods, refrigeration at 4°C and freezing to -20°C, on the determination of BOD, COD and suspended material in water samples. The results were similar and showed that BOD and COD determinations were not affected after 48h storage at 4°C. A significant diminution of these parameters was not observed until after 8 days storage. Freezing at -20°C did not affect COD but caused considerable lowering of BOD, time not being a factor in either case. Storage by both methods caused a significant increase in suspended solids measurements and it is emphasized that they should be carried out immediately after taking samples. 14.3.1.4 Effect of light Shinkar et al. [113] describe a method for the continuous measurement of primary production which is based on the determination of the difference in oxygen concentrations in light and dark conditions. The dissolved oxygen concentration is measured continuously in the sample flowing through two bottles (light and dark), using electrochemical detectors. The readings are transmitted to a multichannel recorder.
858
Oxygen demand parameters
14.3.1.5 Dilution water Nagayama and Hagiwara [114] showed that when the dilution water was allowed to stand in the air incubator at 20°C for 7–10 days, it was possible to make the BOD of the diluent less than 0.2mg L-1. No nitrification was 5 induced with the resulting diluent. 14.3.1.6 Suppression of nitrification Following earlier reports of the use of allylthiourea to suppress the nitrification during BOD determination, Raff [100] examined the suitability of 2-chloro-6-(trichloromethyl)pyridine for the same purpose. The effect of 2-chloro-6-(trichloromethyl)pyridine on BOD was determined for low (less than 25mg L-1) and high (greater than 250mg L1 ) BOD values, for various concentrations of 2-chloro-6(trichloromethyl)pyridine. A 2-chloro-6-(trichloromethyl)pyridine concentration of 50mg L-1 was adequate to suppress nitrification, the effect persisting for more than 14 days as 2-chloror-6(trichloromethyl)pyridine was more resistant to biodegradation than allylthiourea. Concentrations of 200mg L-1 were liable to depress the BOD value attributable to organic compounds. Both 2-chloro-6(trichloromethyl)pyridine and allylthiourea inhibited only the first stage of the nitrification reaction and not the oxidation of nitrite to nitrate. Bird [115] has shown that the extent of nitrification depends on the initial number of nitrifying bacteria associated with suspended particles. Nitrification proceeds more rapidly in the presence of heterotrophic bacteria and suspended solids and the long doubling period of the bacteria and the small number of viable nitrifying bacteria associated with suspended solids result in a small difference between BOD and BOD allylthiourea (ATU) of biological filtration plant effluent. Modifications are recommended for the suppression of nitrification. 14.3.1.7 Kinetics Various workers have investigated the kinetics involved in the BOD test [116–119]. Wilderer et al. [116] used the plateau BOD as a measure of the concentration of biologically degradable substances. A study of the profile of the oxygen consumption curve obtained indicates the presence of a plateau after 8–24h at which the uptake begins to level off before increasing again as a consequence of the endogenous respiration of bacteria, and the contribution from higher organisms and other processes such as nitrification. Measurement of the ‘plateau’ BOD characteristic of the organisms employed can be obtained relatively quickly, the growth curve being approximated by a third-order polynominal equation. The
Biochemical oxygen demand
859
results can be correlated with the BOD value provided a term 5 representing the basal respiration rate is added to the plateau BOD value during the technique described. Barnwell [118] developed a technique, suitable for programmable calculators and small computers, for least-squares estimates of BOD parameters, particularly the rate coefficient. The technique is effective if the BOD time series follows the first-order model and the nitrification stage has been suppressed. The method requires an estimate of the reaction rate coefficient to start an iteration. Successive approximations are found until the difference between them is less than some specified error criterion. The technique has been extended to show how confidence regions for the parameters estimates can be easily determined. A novel algorithm for predicting BOD from the dissolved oxygen 5 concentration after a relatively short incubation period has been described [119]. Tests on synthetic and experimentally derived data indicated that BOD s could be predicted to within 15% in approximately 5 36h. The method was improved by filtering out noise from the raw data via a digital filter. Since the suggested algorithm is simple and does not require a large database, it can easily be implemented on a microcomputer. Middlebrooks [120] has presented a nomogram for solution of the BOD equation. The nomogram solves the ultimate BOD temperature correction, and the rate constant temperature correction. 14.3.2 Sea water It has been reported that whilst the dilution bottle method for BOD yields satisfactory results, on fresh and low saline waters, a discrepancy exists when the test is performed on waters containing elevated levels of sodium chloride and other salts. Seymour [121] found that an increase in salinity resulted in a decrease of BOD values. Degradation rate decreased as the salinity level increased, yet only the particulate fraction seemed to be affected. Degradation of the soluble portion was not affected as adversely by salt content. Lysis of bacteria will occur in a growth medium when sodium chloride is added. It will also occur when salt-acclimatized bacteria are placed in fresh water. If the change in salt concentration is less than 10 000mg L-1 the degree of lysis is negligible, but if greater, the lysis can be extensive. Cellular constituents released following lysis are metabolized by the remaining microbial population in preference to the substrate present, thus yielding erratic BOD results (Kincannon and Gaudy [122, 123]). Davies et al [124] set out to quantify the effects of fresh and saline dilution waters on the BOD of salt water and to quantitatively establish the role of bacterial seed numbers and species associated with changes
860
Oxygen demand parameters
in BOD results. Significant BOD differences were found when saline samples were diluted with standard (non-saline) BOD dilution water. Bacterial populations to genera were monitored and it was shown that equivalent numbers of bacteria did not have the same capability to degrade a given amount of organic matter with increases in salt concentrations to the 3% level. Seeding of hypersaline samples with known salttolerant species is recommended for consistent BOD results. The BOD values obtained in sewage seed organic standards showed a significant trend. BOD values decreased as salt concentrations increased. Dilution of sewage-seeded saline organic standards with standard dilution water resulted in BOD values higher than those of corresponding non-saline organic standards due to increases in bacteria populations and increased organic removal (BOD values) in the presence of low levels of salt. Bacteria populations of sewage seed correlated with corresponding BOD values and concentrations of salt in organic standard solutions show that the addition of 1 and 3% salt resulted in decreased initial bacterial population, decreased growth rates and a decreased ability of an equivalent number of bacteria to degrade an equivalent amount of organic matter. Each salt tolerant bacteria species yielded comparable BOD values for each organic standard conducted at three salt concentrations. Population increased and the rates of biological oxidation were similar at the three salt concentrations when using either the standard dilution water or the saline dilution water. Since sewage seed does not compare favourably with salt-tolerant bacteria in determining the BOD of saline waters, a salt-tolerant bacteria seed should be used to ensure an accurate and reproducible BOD value when saline waste waters are being tested. 14.3.3 Waste waters Several methods for evaluation of waste water strength are available that can be implemented rapidly for both process control and monitoring purposes. Sherrard et al. [125] outline the purposes and requirements of parameters for monitoring and control of waste treatment processes. These authors indicate the limitations of the BOD test and discuss possible alternatives including chemical oxygen demand, total carbon and total organic carbon, total oxygen demand, use of respirometers and comparison between measurement parameters. It is often assumed that measurements of COD, long-term BOD, TOD, the stoichiometric equivalent of TOC, and the long-term respirometric oxygen requirements for a waste water sample should produce equal results. Because each technique is measuring a different combination of waste water properties, however, identical results and absolute oxygen
Biochemical oxygen demand
861
demand correlations between measurement systems cannot be expected. For example, the chemical techniques do not differentiate between readily biodegradable and refractory organic materials. The rapid biological techniques do not describe the long-term oxygen demand that can be exerted by organic material incorporated into aeration basin floc particles. Despite these deficiencies, each measurement provides useful information. The rapid techniques described can aid significantly in effective, efficient, on-line operation of a treatment plant. In addition, their use with batch biological tests can be used to predict plant response to changing influent conditions. Waite [126] evaluated the usefulness of the conventional BOD test and considered possible alternatives or replacements for the BOD determination. The nature of organic contaminants encountered in waste waters, the kinetics of the BOD determination and the relevance of other methods of estimating the organic fraction are discussed, and the nature and methodology of a range of experimental procedures for water quality estimation are reviewed. A single test such as the BOD test is not considered completely satisfactory; the total organic carbon test is considered superior in certain circumstances. Stover and McCartney [127] in the BOD test showed that by making 5 a plot of BOD exerted versus sample volume added to the BOD bottle, the threshold inhibition level of BOD testing can be determined. The threshold inhibition level can be defined as the lowest concentration or volume that causes a reduction in the carbonaceous biological oxidation rate for carbonaceous BOD inhibition. When the BOD of this waste water was measured, with sample values below 3mL of a 1:10 waste water dilution, consistent BOD values were determined. This type of depletion indicating inhibition is often observed in many municipal waste waters and overlooked. It is clear that an understanding of the kinetics and mechanisms of biological growth and respiration is necessary for accurately determining BODs of complex municipal and industrial waste water types. The effects of acclimatized versus non-acclimatized seed material, nitrification, inhibition, dilution seed demand and dilution water depletion must be understood by the analyst conducting BOD testing to minimize variability and obtain reliable results. As a result of a detailed study of the application of the BOD method to waste water samples, Stover and McCartney [127] recommended the following general rules: • •
Use acclimatized seed material with complex waste waters. Inhibit nitrification when appropriate (run both inhibited and uninhibited BOD ). 5
862
• • • • • • •
•
•
Oxygen demand parameters
Determine seed demand and dilution water depletions with multiple dilutions. Apply appropriate correction factors. Dilution water correction can be calculated according to the amount of dilution water used if desired. Generally, if greater than 200mL of dilution water is used, the total water correction can be used. Select seed concentration to yield between 0.2 and 1.0mg L-1 oxygen demand. During data interpretation, place greater emphasis on sample volumes with higher total depletion. The influences of water and seed correlations are minimized during the calculation of these BOD values. Best sample volumes for BOD calculations are those with net depletions between 25 and 75% of the initial dissolved oxygen (DO) values. Place more emphasis on sample volumes with greater than 2.0mg L1 net depletion. Do not use sample volumes with final DO values below 1.0mg L-1. Plot BOD versus sample volume to determine inhibition. Use sample volumes below the threshold inhibition level for BOD calculations.
Although the BOD test is run as a series of dilutions, the result from each dilution is ordinarily analysed separately from the rest of the series of dilutions. By combining the results observed from each dilution, using appropriate data points and considering the effects of the previously described factor of BOD testing, the variability of BOD testing can be minimized. Tests on waste water samples have shown [128] that intermittent stirring tended to increase the BOD value by about 7%. This increase is 5 not approximately the same order of magnitude as the error inherent in the BOD determination itself and it is therefore concluded that for routine determinations, stirring is unnecessary. Ademoroti [129] studied the effects of copper, zinc, sulphate, nitrate and chloride contaminants on the BOD of waste waters. One microgram per litre of copper or zinc, respectively, suppressed BOD by 33 and 16.9%. Parisod and Schroeder [130] carried out experiments with glucose, glutamic acid, succinic acid and sorbitol to determine whether plateau BOD values obtained from mixed substrate systems were stoichiometric end points for the substrate conversion reactions and to investigate the kinetics of the mixed substrate system. It was found that the plateau values resulting from substrate mixtures could be closely predicted by adding the average plateau values for the individual components. It was also shown that the ultimate BOD of a waste water can be determined
Biochemical oxygen demand
863
by adding the plateau BOD and the theoretical BOD of the cells produced up to the plateau. 14.3.3.1 Microbial electrodes Hirata et al. [131] developed equipment for the rapid determination of BOD by means of a microbial electrode of immobilized living whole cells of yeast, a porous membrane and an oxygen electrode. The oxygen electrode current indicated the level of oxygen saturated when air was bubbled through a water sample containing no organic matter; the current decreased when this water was replaced with a sample containing organic matter. Tests were carried out on various domestic and industrial waste waters. Good correlations were obtained between BOD values determined by this technique and those determined by the conventional BOD method. 5
14.3.3.2 Respirometric methods Respirometers can be used to determine the oxygen demand of a waste water exerted by a specific biological culture. The test is relatively easy to perform and preliminary data for monitoring purposes can be available in less than 3h. If used with rapidly settled influent waters and fresh aeration tank solids, respirometric data can be used to predict aeration tank culture responses to changing oxygen demands. Commercially available respirometer systems are easily used and inexpensive to operate. Respirometric techniques have the advantage of providing operators with toxicity information when correlated with non-biological tests. Damaschke [132] has described an apparatus for the automatic determination of BOD in undiluted waste water by the continuous measurement of the oxygen consumption. The oxygen consumption is recorded continuously for six measuring positions simultaneously and a BOD time curve is plotted, from which the 5-day oxygen consumptions can be evaluated. Tuffy et al. [133] carried out a critical analysis of Warburg respirometry for BOD determinations on waste waters. The reliability of the Warburg technique for estimation of BOD at levels of 10mg L-1 or less was examined using water samples from a stream receiving discharges of primarily domestic origin. The results indicate that the Warburg method can be used to determine low levels of carbonaceous BOD but that the variance is an order of magnitude greater than that of the alternative technique employing a BOD bottle and a dissolved oxygen probe. The method is, however, unsuitable for nitrification studies since, although ammonia is oxidized to nitrite under the
864
Oxygen demand parameters
experimental conditions, the action of Nitrobacter, and the consequent oxidation of nitrite to nitrate, is inhibited by the potassium hydroxide used to absorb carbon dioxide. Tuffy et al. [133] use Amino Warburg respirometers with 125mL flasks and 50mL sample volumes; 1mL of 15% potassium hydroxide was placed in the centre well as a carbon dioxide absorbent. Temperature was maintained at 20°C for the duration of the runs, usually 15–20 days. Standard operating procedure consisted of equilibrating the sample to 20°C and aerating it to saturation. Manometer readings were recorded to the nearest millimetre and incubation time to the nearest 0.01 days. In order to determine the variances of the conventional and Warburg BOD techniques, nitrification inhibited samples, uninhibited samples, and thermobarometers were incubated. Inhibition was achieved with 0.5mg L-1 of allylthiourea replenished at 7-day intervals. Inhibited and uninhibited samples were also incubated at 20°C with the BOD bottle technique. Standard 200mL BOD bottles were used and dissolved oxygen was recorded to the nearest 0.02mg L-1. The study on nitrification was accomplished by comparing the oxygen demand, change in nitrogen species, and the enumeration of Nitrosamones and Nitrobacter found in the Warburg sample to that found in a control. In nitrification, the ammoniacal nitrogen in the system undergoes oxidation by the bacterial species Nitrosamonas to nitrite and then oxidation from nitrite to nitrate by the species Nitrobacter. These reactions proceed according to the following equations respectively: NH + 3/2O → HNO + H O
(14.1)
HNO + 1/2O → HNO
(14.2)
3
2
2
2
2
2
3
The associated oxygen consumption would be 3.43mg oxygen for every 1.0mg of nitrogen oxidized in eqn (14.1) and 1.14mg oxygen for every 1.0mg of nitrogen oxidized in eqn (14.2). Experimental results [134] indicate a slightly lower combined oxygen demand of 4.3mg L-1 per 1mg L-1 of ammoniacal nitrogen. Figure 14.1 is a plot of the carbonaceous BOD data as well as total combined BOD obtained by the two techniques. The differences between the total combined and the carbonaceous BOD is that portion of the BOD due to nitrification. The nitrogenous fraction calculated from the BOD bottle dissolved oxygen (DO) probe technique is 34.22mg L-1, whilst that from the Warburg data is only 20.33mg L -1 . Some of this difference is due to inaccuracies in
Biochemical oxygen demand
865
Fig. 14.1 BOD of Papssaic River sample with and without nitrification as determined by Warburg and BOD bottle DO probe technique. ( ) with nitrification, BOD-DO; (X) with nitrification, Warburg; (°) with nitrification, BOD-DO; (∆) without nitrification, Warburg.
.
measuring the 20-day carbonaceous BOD but the greatest portion is accounted for in an interference to the normal course of nitrification by the Warburg technique. The oxygen respirometric method of measuring the oxygen uptake of waste waters has also been discussed by Clark et al. [135]. Schoenen and Thofern [136] using samples of synthetic waste water made measurements of BOD using an automatic manometric apparatus under various conditions, to investigate factors affecting the results, such as the speed of stirring, size of magnetic stirrer and shape of the measuring flask. Their results indicate the importance of ensuring constant, optimal and reproducible conditions for BOD tests. It is especially indicated that an optimal input of oxygen should be ensured, by a high rate of stirring, to avoid adverse effects on the microbial growth; typical results are given showing the advantages of increasing the rate of stirring from about 350 to 700rev min-1. The use of oxygen demand curves to asses the toxic effects of a waste water is also considered.
866
Oxygen demand parameters
14.3.3.3 Correlations between oxygen demand parameters Work on the correlations of oxygen demand parameters measured in waste waters, principally COD, BOD and TOD, is summarized in Table 14.4 [140–147, 156, 157]. A fair assessment of the situation is now seen regarding these correlations and is as follows. Analysis of total organic carbon in plant effluents, receiving waters, and in water and waste water treatment processes is rapidly gaining favour as the most reliable index of waste loading and stream pollution. If the waste is from a single source, or relatively constant in nature, reasonably good correlation is usually found between TOC, BOD and COD. If the waste is highly variable in composition, correlation among the methods is lost. This is not too surprising because there is no inherent correlation between BOD and COD. The COD method does not oxidize certain organics which are measured in the TOC procedure and are biodegraded in the BOD test. Results in both are affected by certain oxidizable inorganic compounds, while TOC measurement is unaffected by those inorganic compounds in concentrations normally found in the waters to be analysed. 14.3.4 Trade effluents Various workers have discussed particular aspects of the standard BOD method in relation to measurements made on trade effluents 5 (Table 14.5). A departure from the classical method for determining BOD is the electrolysis method described by Young and Baumann [139]. In this procedure the oxygen content of the space about the sample is enriched with electrolytically generated oxygen. As the oxygen is used up by the sample the concentration in the headspace is made up and consequently the amount of oxygen reacted is obtained. Table 14.4 Correlation of oxygen demand parameters of waste waters
Biochemical oxygen demand
867
Table 14.5 Determination of biochemical oxygen demand of trade effluents
Young and Baumann [139] conducted BOD measurements under air containing 21% oxygen and under an atmosphere of about 80% oxygen and obtained good agreement between the results. Chemical methods of controlling nitrification were used in both methods. The electrolysis BOD method provides a direct and continuous measure of oxygen uptake in an effluent sample. The principal advantages of this method are that (1) little or no dilution of the sample is required, (2) larger and more representative samples can be analysed, (3) tests can be run for days or weeks without interruption, (4) the equipment is simple to operate, (5) no titrations are required and oxygen uptake can be obtained by simply reading a dial and (6) BOD can be measured with significantly greater precision than by the classical method. BODs obtained by electrolysis measured at any time after 2 days of incubation can be correlated reasonably well with those obtained by standard dilution. The most promising use of the method however, was not for a single point measurement but for evaluating the oxygen uptake by characteristics of effluents and for determining the effect of dilution and the presence of toxic materials on effluent treatability. 14.3.4.1 Relationship between oxygen demand parameters Voorn [148] has reported that he obtained an excellent correlation between COD and TOD on settled samples of industrial effluents. Arin [149], on the other hand, found that in the case of effluents containing suspended matter the TOD measurement was consistently
868
Oxygen demand parameters
greater than the COD but both followed the same trend. The discrepancy between the two measurements was attributed to the more efficient oxidation of the particulates in the TOD measurement. 14.3.5 Sewage effluents Extensive work has been carried out on various aspects of the measurements of the BOD of sewage effluents. This is reviewed in Table 14.6. Topics covered include inhibition of nitrification during measurements on sewage [150–153], effect on BOD of toxic substances [154, 155], sample preservation [160], and reaction kinetics [157–159]. The relationship between BOD and other oxygen demand parameters has also been discussed [160–164]. More recent work has been concerned with the semi or full automation of the BOD procedure. Crowther et al. [102] have described a semiautomated procedure which has been applied to sewage and other types of samples. This method was capable of handling 1000 samples per week. Galovatyi and Smirnov [165] described an instrumental method of measuring the rate of consumption of oxygen for monitoring the biochemical demand of unpurified sewage effluents. The method is based on measuring the rate of oxygen consumption by a mixture of activated sludge and effluent and a reference liquid in a flow-through respirometer. The latter comprises preparation vessels, metering pumps, pulse generator, recorder, current stabilizer, electrolyser, pressure sensor, over-flow device and fermenter. The method has an accuracy of 20% and is suitable for routine BOD monitoring of unpurified effluents and for use in municipal sewage works. Reigler [166] has described a continuous BOD measuring device. The equipment continuously samples a sewage or effluent flowing through a coarse cylindrical screen inserted into a pipeline, measures the oxygen consumed by the suspended biomass and prints out a continuous BOD curve. Owing to the short measurement interval of 3min, a continuous record of fluctuations in the strength of incoming or outgoing sewage is possible and a measurement range from 5 to 500mg L -1 can be accommodated. The screen retains particles greater than 0.5mm in diameter, its surface being self-flushing. The liquid passing through is diluted with tap water and the oxygen uptake measured by oxygenselective electrodes at the inlet and outlet from a culture flask in which the biomass is supported on a number of light plastic rings. The dilution ratio is selected to maintain the rate of oxygen uptake in the suspended growth medium constant at 5mg oxygen per litre. Damiecki [168] has stated that BOD5 determinations should be performed in the presence of 0.5mg L-1 of allylthiourea as a nitrification
Biochemical oxygen demand
869
Table 14.6 Measurement of biochemical oxygen demand of sewage
inhibitor. A comparison of BOD values of effluents from biological treatment plants with a wide range of sludge and volumetric loadings, with and without the addition of allylthiourea is reported. Both for biological filters and for activated sludge plants the values determined without the addition of allylthiourea were markedly higher than those for which nitrification was inhibited. The difference decreased from over 60% at low values of organic sludge loadings (200 BOD m-3 per day) to about 20% at 750 BOD m-3 per day and over. Washington et al. [169] determined the amount of nitrogenous oxygen demand as measured in the standard BOD tests associated with 5 different types of sewage waters from various sources. It was found that a substantial BOD was exerted in both completely and partially nitrified sewage and primary and secondary effluents, including chlorinated
870
Oxygen demand parameters
effluent. Alternative techniques for limiting nitrogenous oxygen demand, to comply with effluent BOD standards are indicated. 14.3.5.1 Microbial electrodes Karube et al. [170] and Strand and Carlson [171] have discussed microbial electrode BOD sensors. Karube et al. [170] evaluated two types of sensor for estimating BOD. The first consisted of a bacteria collagen membrane and oxygen electrode. The current of the electrode decreased to a steady state which had a linear relationship with the BOD of solution when the electrode was inserted in a solution of glucose and glutamic acid. BOD can be estimated in 15min. The second sensor was a biofuel cell using an immobilized Clostridium butyricum platinum electrode. The steady-state current was attained in 30–40min at 37°C and was proportional to BOD. The relative error of the BOD estimation was within ±10%, and the current output of the biofuel cell was almost constant for 30 days. Strand and Carlson [171] carried out rapid BOD measurements on municipal waste waters using a biofilm electrode. They compared the efficiency of a biofilm electrode with the standard BOD test for samples of crude, settled and biologically treated sewage. Two immobilized membranes (biofilms) were prepared by applying centrifuged cell paste to a nylon net; these were placed on the surface of a dissolved-oxygen electrode covered with a Teflon membrane and the biofilm was covered with a polycarbonate membrane filter and a protective layer of nylon mesh. The biofilm probe gave a signal within 20min which was proportional to the conventional BOD for 5 soluble organic matter in settled and treated sewage but did not respond to the particulate organic matter in crude sewage. The precision of the results obtained was comparable to that of the conventional BOD test and the electrode remained stable for up to 3 5 weeks. 14.3.5.2 Kinetics Ogunrombi and Onuoha [172] present a second-order model of the kinetics of the BOD reaction. Composite samples and crude sewage waste water from the inlet to the sewage treatment plant, were analysed for BOD and COD; hydraulic flows were estimated by using a throated flume. Values of the deoxygenation rate constant were estimated on the basis of the second-order kinetics. The waste water was of medium strength, with an average BOD of 310mg L-1 and COD 5 of 491mg L -1 , and an approximately linear relationship existed between BOD and COD.
References
871
References 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41
Korenaga, T. and Ikatsu, H. Analyst (London) 106 653 (1981) Viraraghavan, T. Water Sew. Works 123 80 (1976) Golterman, H.L. and Wisselo, A.G. Hydrobiologia 77 37 (1981) Dugin, G.V., Pisarevskii, A.M. and Polozova, I.P. Soviet J. Water Chem. Technol. 7 69 (1985) Lur’e, Y.Y. and Nikolaeva, Z.V. Zhur. Analit. Khim. 28 1433 (1973) Andrews, C. and Schierjott, G. Water Waste Engng. 11 45 (1974) Baughman, G.L., Butler, B.T. and Sanders, W.M. Water Sewage Works 116 359 (1969) Kelkenberg, H. Z. Wasser Abwasser Forsch. 8 146 (1975) Zeitz, W. Wasserf. Wasser Abwasser 117 181 (1976) Wagner, R. and Ruck, W. Z. Wasser Abwasser Forsch. 14 145 (1981) Wagner, R. and Ruck, W. Z. Wasser Abwasser Forsch. 15 287 (1982) Messenger, A.L. Journal of Water Pollution Control Federation 53 232 (1981) Ryding, S.O. and Forsberg, A. Water Research 11 801 (1977) Aslam, S. and Walker, O.L. Water Pollut. Contr. Fed. 54 1148 (1982) Gudernatsch, H. and Doppelfeld, W. Z. Wasser Abwasser Forsch. 8 182 (1975) Moore, L. J. Water Pollut. Contr. Fed. 48 2327 (1976) Clark, D.W. and McKinney, W.M. Water Wastes Engng 14 45 (1977) Watter, R.J. and Adams, V.D. Water Research 17 715 (1983) Ogneau, R.C. Hidrotechnica 33 195 (1988) Le Clair, G., Des Jardins, F., Briere, F., Millette, L. and Dagenais, D. Sci. Techn. L’eau 20 131 (1987) Wolf, G. and Oliver, M. Umwelt No. 5 512 (1980) Fleet, B., Ho, A.Y.W. and Tenygl, J. Analytical Chemistry 44 2156 (1972) Fleet, B. Talanta 19 317 (1972) Te Kippe, R.J. and Zehnefenning, R.G. J. Water Pollut Contr. Fed. 43 515 (1971) Phillips, J. Water Research 3 911 (1969) Phillips, J. Water Research 3 907 (1969) Chaudhuri, N., Nujogi, S. De A. and Basu, A. J. Water Pollut. Contr. Fed. 45 537 (1973) Warzee, P. Foundation of the University of Luxembourg, Arlon, Belgium. Research Notes No. 13 ( P13 WAR), Automatic Determination of OCD of Water Samples (1977) Thomas, O. and Muningda, Y. Trechn. Sci. Munic. 75 277 (1980) Southway, C. and Bark, L.S. Anal. Proc. (London) 18 7 (1981) Takimoto, K., Kuratoni, H., Eto, A., Mukai, T. and Komuro, M. Water Wastes 22 187 (1980) Ballinger, D., Lloyd, A. and Morrish, A. Analyst (London) 107 1047 (1982) Fleet, B., Ho, A.Y.W. and Tenygi, J. Analyst (London) 97 321 (1972) Akesson, M. and Lind, J.E. Valten 39 217 (1983) Department of the Environment and National Water Council, Biochemical Oxygen Demand: Examination of Waters and Associated Materials, HM Stationery Office, London (1981) Kempa, E.S. Oesterreichische Abwasser – Rundschau 21 20 (1976) Lloyd, A. Analyst (London) 107 1316 (1982) Jirka, A.M. and Carter, M.J. Analytical Chemistry 47 1397 (1975) Canelli, E., Mitchell, D.G. and Pause, R. Water Research 10 351 (1976) Korenga, T. and Ikatsu, H. Anal. Chim. Acta 141 301 (1982) Shriver, L.E. and Young, J.C. J. Water Pollution Control 44 2140 (1972)
872
Oxygen demand parameters
42 Appleton, J.M.H., Tyson, F. and Mounce, R.P. Anal. Chim. Acta 179 269 (1986) 43 Pokorny, S. Papir. Cellulose 27 3 (1972) 44 Wells, W.N. Water Sewage Works 117 123 (1970) 45 Bonnefille, M. and Beauchamp, R. Tech. Sci. Munic. 73 247 (1978) 46 Leoni, C, Lovato, O.G. and Belluci, G. Industria Conserve. 48 19 (1973) 47 British Standards Institution (London) BS2690 Part 13. Methods of testing water used in industry, part 13, Dichromate value (chemical oxygen demand) non-volatile organic carbon, tannins and chlorine (1972) 48 Leith, W. Chemikerzeitung – Chem. Appar. 95 463 (1971) 49 Edwards, S.J. and Allen, M. Analyst (London) 109 671 (1984) 50 Fisher, D.B. Water Pollut. Contr. 113 24 (1979) 51 Frieg, H. Z. Wasser Abwasser Forsch. 15 266 (1982) 52 Smith, R. Water South Africa 6 204 (1980) 53 Ruider, E. and Spatzierer, G. Abwassertechnik 32 10 (1981) 54 Morfaux, J.N. and Sarris, J. Trib. Cebedeau 27 97 (1974) 55 Hey, A.E., Green, A. and Harkness, N. Water Research 3 873 (1969) 56 Malissa, H. and Bleier, H. Mitt. Geb. Lebenssmittel, Hyg. 62 382 (1971) 57 Merz, W. Z. Wasser Abwasser Forsch. 11 171 (1978) 58 Stenger, V.A. and Van Hall, C.E. J. Water Pollut. Contr. Fed. 40 1755 (1968) 59 Carr, J.D. US National Technical Information Service, Springfield, Virginia. Report no. PB 271439. Use of potassium ferrate in oxygen demand measurement (1977) 60 Bontin, P., Bechac, J.P., Don, J.L., Mercier, B. and Roy, J.L. Tech. Sci. Munic. 22 239 (1977) 61 Haapala, K. and Starck, B. Valten 42 311 (1986) 62 Corning, D.R. BLCJ 31 101 (1988) 63 Clark, D.W., Mitchell, R.E., Archuleta, A. and Barkley, W. Water Sewage Works 122 94 (1975) 64 Virarajhavan, T. J. Water Pollut. Contl. Fed. 48 2213 (1976) 65 Fornsbery, A. and Ryding, S.V. Valten 31 148 (1975) 66 Wolf, P. and Nordmann, W. Correspond. Abwasser 24 277 (1977) 67 Schoke, K. Korrespond. Abwasser 28 275 (1981) 68 De Casseres, K.E., Best, D.C. and May, B.D. Water Pollut. Contr. 83 416 (1984) 69 De Casseres, K.E. Water Pollut. Contr. 79 143 (1980) 70 Best, D.G. and De Casseres, K.E. Water Pollut. Contr. 77 138 (1978) 71 Stones, T. Effl. Water Treat. J. 18 290 (1978) 72 Goodwin, A.E., Cabbiness, D.E. and Mottola, H.A. Water, Soil Pollut. 8 467 (1977) 73 Musselwhite, C.C. and Lowden, G.F. Water Research Centre Technical Memorandum TM 118. Evaluation of the Wosthoff Hydromat. COD Analyser, February (1976) 74 Voorn, G. and Marlow, J.S. Water Treatment Journal 302, June (1977) 75 Von Becker, W.J. Z. Wasser Abwasser Forsch. 12 196 (1979) 76 Voorn, G. and Marlow, J.S. Effl. Water Treat. J. 17 301 (1977) 77 Voorn, G. and Frampton, J.L. J. Water Pollut. Contr. Fed. 114 32 (1976) 78 Voorn, G. and Mitchell, P. Int. Labor. 25 Jan–Feb (1976) 79 Ravenscroft, B.D. Paper presented to the Society for Analytical Chemistry, North East and Scottish Regions, and the Society for Water Treatment and Examination, Newcastle, UK, February (1975) 80 Briggs, R., Schofield, J.W. and Gorton, P.A. J. Water Pollut. Contr. Fed. 114 47 (1976)
References
81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97
98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116
873
Laffont, C, Capdeville, B. and Rogues, H. Water Research 13 167 (1979) Capdeville, B. and Rogues, H. Water Research 14 501 (1980) Clifford, D.A. and McGaughey, L.M. Analytical Chemistry 54 1345 (1982) Fukunaga, I., Oda, K., Lida, S. and Uno, G. Water Purif. Liqu. Wastes Treat. 20 509 (1979) Nakajima, K., Ishizuka, T. and Sunahara, H. Water Research 16 1003 (1982) Legrand, C. Capdeville, B., Rogues, H. and Angles, M. Int. J. Environ. Anal. Chem. 11 283 (1982) Arin, M.L. Environ. Sci. Technol. 8 898 (1974) Briggs, R., Schofield, J.W. and Gorton, P.A. Water Pollut. Contr. 75 47 (1976) Ekehahl, G. Valten 32 328 (1976) Marty, J.L. and Aim, R.B. Tech. Sci. Munic. 71 531 (1976) Wells, W.N. Water Pollution Control Federation, Deeds and Data D2-D3, February (1972) Ravenscroft, B.D. Pollut. Water Chem. 254 February (1975) Voorn, G. and Frampton, O. Water Waste Treatment 20 10 (1977) Stones, T. Water Pollut. Contr. 73 673 (1974) Cummings, J.P. British Patent 1,314,917 (12/7/71) Musselwhite, C.C. Water Research Centre, Medmenham, UK Technical Memorandum TM 107. A limited evaluation of total oxygen demand meter (1975) Department of the Environment/National Water Council Standing Committee of Analysts. Methods for the Examination of Waters and Associated Materials. Biochemical oxygen demand, 1981 , HM Stationery Office, London (1983) Hewitt, J., Hunter, J.V. and Lockwood, D. Water Research 13 325 (1979) Klein, R. and Gibbs, C.R. Journal of Water Pollution Control Federation 51 2257 (1979) Raff, J. Gas- u. Wasserf. Wasser Abwasser 122 301 (1981) Rigin, V.I., Golovin, Y.G. and Tyuneva, G.S. Soviet Journal of Water Chemistry and Technology 4 113 (1982) Crowther, J.M., Dalrymple, J.F., Woodhead, T., Coackley, P. and Hamilton, J. Water Pollut. Contr. 77 525 (1978) Friege, H. and Bohme, A. Umwelt No. 4 236 (1982) Busse, H.J. Z. Wasser Abwasser Forsch. 8 164 (1975) Gantner, H. Fresenius Z. für Analytische Chemie 299 42 (1979) Kohne, M., Sicpmann, F.W. and Te Heesen, D. Korrespond. Abwasser 33 787 (1986) Strigel, G. Gas- u. Wasserf. Wasser Abwasser 120 216 (1979) Steinecke, H. Gas- u. Wasserf. Wasser Abwasser 117 454 (1976) Steinecke, H. Gas- u. Wasserf. Wasser Abwasser 120 21 (1979) Steinecke, H. Gas- u. Wasserf. Wasser Abwasser 120 50 (1979) Ranchet, J., Pescheux, F. and Menissier, F. Bull. Liaison Laborator. Pont Chausers 106 27 (1980) Ranchet, J., Pescheux, F. and Menissier, F. Techn. Sci. Munic. 76 547 (1981) Shinkar, G.G., Sapozhnikov, V.V. and Georgiev, A.A. Water Resources 8 205 (1981) Nagayama, T. and Hagiwara, K. Water Purif. Lyn. Waste Treatment 20 323 (1979) Bird, P.G. Water Pollution Control 80 378 (1981) Wilderer, P., Harman, L. and Keser, G. Z. Wasser Abwasser Forsch. 11 82 (1978)
874
Oxygen demand parameters
117 118 119 120
Marske, D.M. and Polkowski, L.B. J. Water Pollut. Contr. Fed. 44 1987 (1972) Barnwell, T.U. J. Environ. Engng Div. ASCE, 106 1197 (1980) Raviv, R. and Ben-Yaakov, S. Biotechnol. Bioengng. 26 1239 (1984) Middlebrooks, E.J. Water and sewage Works Ref. F 153 and R 155–156 (1975) Seymour, M.P. The biodegradation of particulate organic matter in seawater media, Thesis, Rice University (1977) Kincannon, D. R and Gaudy, A.F. Biotech. Engng. 8 371 (1966) Kincannon, D.F. and Gaudy, A. J. Water Pollut. Contr. Fed. 38 1148 (1966) Davies, B.M., Bishop, J.R., Gurhrie, R.K. and Forthafers, R. Water Research 12 917 (1978) Sherrard, J.H., Friedman, A.A. and Rand, M.C. J. Water Pollut. Contr. Fed. 51 1799 (1979) Waite, T.D. Australian Water Resources Council, Canberra, Technical Paper No. 34. An evaluation of non-specific measurements on organic pollutants in waters and waste waters (1978) Stover, E.L. and McCartney, D.E. Environmental Engineering Consultation Inc. Stillwater, Oklahoma, Private communication McGhee, T.J. and Walsh, R.D. Water Sewage Works 121 46 (1974) Ademoroti, C.M.A. Biolog. Wastes 24 259 (1988) Parisod, J.P. and Schroeder, E.D. Journal of Water Pollution Control Federation 50 1827 (1978) Hirata, K., Otani, Y., Hikuma, M. and Yasuda, T. Workshop Houston and Denver, 529 BOD quick estimating system utilizing a microbial electrode (1985) Damaschke, K. Gesundheits-Ingenieur 93 245 (1972) Tuffy, T.J., Hunter, J.V. and Hewitt, J.P. Proceedings of the 29th Industrial Waste Conference, Purdue University, US A. A critical analysis of working respirometry for BOD determinations on polluted water (1974) Wezernak, C.J. and Cannon, J.J. J. Sanit. Eng. Div. ASCE 94 883 (1968) Clark, A.N., Eckenfelder, W.W., McMullen, E.D., Roth, J.A. and Young, B.A. Water Research 12 799 (1978) Schoenen, D. and Thofern, E. Gesundheits-Ingenieur 96 329 (1975) Eggink, H.J. H2O 10 282 (1977) Sybrandi, J.C. H2O 9 274 (1976) Young, J.C. and Baumann, E.R. Engineering Research Institute, Iowa State University, Ames, Iowa. Demonstration of the Electrolysis method for determining BOD (1972) Stones, T. Water Pollut. Contr. 71 431 (1972) Ajmial, M., Ahmed, A. and Nomani, A.A. Water Research 16 1611 (1982) Casser, P.R. Eff. Water Treat. J. 22 357 (1982) Hikuma, M., Suzuki, H., Yasuda, T., Karuba, C. and Suzuki, S. Europ. J. Appl. Microbiol. Biotechnol 8 289 (1979) Wagner, R. Gas- u. Wasserfach Wasser – Abwasser 117 443 (1976) Hiidenheimo, H. and Wilson, M.F. Water Research 8 535 (1974) Hwang, C.P., Goos, G. and Davis, E. Water Pollut. Contr. 112 28 (1974) Douglas, T.B. Tappi. 61 53 (1978) Voorn, G. Sci. Ind. No. 11 10 (1978) Arin, L.A. Environ. Sci. Technol. 8 901 (1974) Hjart, J., Bengstom, G., Ekedahl, G. and Lindgren, O. Valten 41 259 (1985) Dague, R.E. J. Water Pollut. Contr. Fed. 53 1738 (1981) Hall, J.C. and Foxen, R.J. J. Water Pollut. Contr. Fed. 55 1461 (1983)
121 122 123 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152
References
875
153 Stamer, J.K., McKenzie, S.W., Cherry, R.N., Scott, T.C. and Stamer, S.L. Water Pollut. Contr. Fed. 51 918 (1979) 154 Berkun, M. Water Research 16 559 (1982) 155 Damascke, K. Gesundheits Ingenieur 95 309 (1974) 156 Neis, U. Z. Wasser Abwasser Forsch. 11 3 (1978) 157 Hewitt, J.P. National Technical Information Service, Springfield, Virginia, PB-268518. A comparison of methods for calculating constant for the first order BOD equation (1971) 158 Sletten, O. Proceedings of the 26th Industrial Waste Water Conference, Purdue University. Methods for operators of waste water treatment plants to determine BOD parameters, p. 740 (1971) 159 Stones, T. Water Pollut. Contr. 38 435 (1984) 160 Wilderer, P., Engelman, G. and Schmenger, H. Gas- u. Wassfach. Wasser Abwasser 118 357 (1977) 161 Stones, T. Water Pollut. Contr. 80 659 (1981) 162 Clark, D.W. Water Sewage Works 121 68 (1974) 163 Constable, T.W. and McBean, E.R. Water Air Soil Pollut. 11 363 (1979) 164 Reynolds, J.F. and Goellner, K.A. Water Sewage Works 121 31 (1974) 165 Galovatyi, E.I. and Smirnov, D.N. Soviet J. Water Chem. Technol. 5 99 (1983) 166 Reigler, G. Korrespond. Abwasser 31 369 (1984) 167 Brown, J.A., Paul, J., Rice, L.C. and Lines, J.M. Water Sewage Works 120 105 (1973) 168 Damiecki, R. Korrespond. Abwasser 27 832 (1980) 169 Washington, B., Luc-Hing, C, Zenz, D.R., Rao, K.C. and Obayashi, A.W. Water Pollut. Contr. Fed. 55 1196 (1983) 170 Karube, L., Matsunaga, T., Mitsuda, S. and Suzuki, S. Biotechnol. Bioengng. 19 1535 (1977) 171 Strand, S.E. and Carlson, D.A. J. Water Pollut. Contr. Fed. 56 464 (1984) 172 Ogunrombi, J.A. and Onuoha, J.C. Ind. J. of Environmental Health 24 219 (1982)
Index
Abate, determination of 569–570 Adenosine triphosphate, determination of 478–481 Alcohols, determination of 248–250 Aldehydes determination of 250–253 preconcentration of 251–252 Aldrin, determination of 517 Aliphatic amines, determination of 434–435 Aliphatic hydrocarbons determination of 109–140 preconcentration of 129– 136 5-Alkyhydantoin, determination of 470 Amides, determination of 444–447 Amino acids determination of 453 preconcentration of 455 Aminophenols, determination of 440 Aminotriazole, determination of 470 Anionic detergents determination of 295–315 preconcentration of 312, 314 Anodic scanning voltammetry, determination of organocadmium compounds 770 Aromatic amines determination of 437–439 preconcentration of 438 Aromatic hydrocarbons determination of 140–144 preconcentration of 144 Aryl phosphates, determination of 477–478 Atomic absorption spectrometry, general 15–17 Atomic absorption spectrometry of anionic detergents 305 carboxylic acids 242 cationic detergents 317 ethylene diamine tetracetic acid 460 metals in oil 197 nitrilacetic acid 468 non-ionic detergents 327, 328, 332,
333 organoantimony compounds 709 organoarsenic compounds 725– 727 organolead compounds 731, 733 organomercury compounds 735– 741, 745, 747–755 organotin compounds 762, 767 phenols 277 Atomic emission spectrometry, general 24 Atomic emission spectrometry, determination of germanium 769 Azarine, determination of 470 Bayrusil, determination of 571 Benethionium salts, determination of 487 Benzoic acid, determination of 233 Benzothiazole, determination of 488, 489 Biochemical oxygen demand automation of 856 determination of 854–870 effect of experimental parameters on 855–859 microbial electrodes 863–870 respirometry 863–865 Bromine, determination of 783, 784 Calcium lignosulphonate, determination of 493 e-Caprolactam, determination of 470 Carbamate insecticides and herbicides determination of 258–261 preconcentration of 582 Carboxylic acids determination of 232– 248 preconcentration of 241 Carboxymethyl succinate, determination of 630–631 Cationic detergents, determination of 315–318
Index
Chemical oxygen demand, determination of 838–870 Chemiluminescence analysis, determination of organic nitrogen 796 Chemiresist detector, determination of organophosphorus compounds 482 Chlorinated alkyl naphthylenes, determination of 395 Chlorinated insecticides determination of 496–542 preconcentration of 514, 515, 522, 533, 534 Chlorinated isocyanurates, determination of 420, 470 Chlorinated polyaromatic compounds, determination of 395 Chlorine, determination of 778–831 Chloroamines, determination of 470 Chloroamino acids, determination of 455 Chloroanisoles, determination of 422 Chloroaromatic compounds, determination of 439, 440 Chlorolignosulphonic acids, determination of 420, 493 Chloromequat, determination of 608 Chloromethyl anisoles, determination of 422 Chloromethyl ethers, determination of 422 Chloromethylsulphenyl bromobenzene, determination of 604 Chlorophenols determination of 407, 417 preconcentration of 412, 417 Chlorophyll, determination of 648–660 Cobalamin, determination of 638, 639 Column chromatography, general 70– 71 Column chromatography of aliphatic amines 435 anionic detergents 312 aromatic hydrocarbons 142, 158 azarenes 470 carboxylic acids 248 chloroaromatic amines 439 cobalamin 638, 639 ethylenediamine tetraacetic acid 459–461 haloaromatic compounds 395 nitriloacetic acid 467, 468 nitrophenols 443 non-ionic detergents 322–325 organophosphorus insecticides 563 ozonization products 633–638
877
Coprostanol, determination of 628, 629 Coral, determination of 571 Coumaphos, determination of 571 Dalapon, determination of 607, 608 DDT, determination of 516, 517 5,5-Dialkylhydantoin, determination of 470 Diazo compounds, determination of 470 Dicamba, determination of 601 Dichlorobenil, persistance of 601 2,3-Dichloro-1,4-naphthaquinone, determination of 631 Dichlorvos, determination of 515, 516 Dieldrin, determination of 518 Dihydroxyabietic acid, determination of 633 S-2(Diisopropylamino)ethyl-Oethylmethylphosphonothioate, determination of 631 Dimethyl sulphide, determination of 489–492 Dioxans, determination of 262 Dioxins, determination of 421, 422 Diquat, determination of 604, 605 Dissolved organic carbon, determination of 826–830 Disulphides, determination of 484–487 Dodecyl guanidines, determination of 470 Dursban, determination of 568, 569 Emission spectrometry, determination of organosulphur compounds 806 Endosulphan, determination of 517, 518 Endrin, determination of 518 Enzymic assay, general 33 Enzymic assay of carbamate insecticides/herbicides 581 carbohydrates 258 chlorinated insecticides 513 cobalamin 638, 639 S-2(diisopropylamino)ethyl-Oethylmethylphosphonothioate 631 isopropylmethylphosphonofluoridate 631 organophosphorus insecticides 564, 565 phenols 285 polychlorobiphenyls 402 triazine herbicides 592 Esters, determination of 253–258 Ethers, determination of 289
878
Index
Ethylenediamine tetraacetic acid, determination of 458–462 Ethynyloestradiol, determination of 630 Fenitrothion, 570, 571 Flow injection analysis, general 13–15 Flow injection analysis, determination of amides 446 cationic detergents 316, 317 chemical oxygen demand 844 non-ionic detergents 333, 334 organic nitrogen 797 organophosphorus insecticides 543 Fluorescence spectroscopy, general 7– 11 Fluorescence spectroscopy of amino acids 453, 455 Bayrusil 571 carbamate insecticides/herbicides 581 carboxylic acids 240 chlorophyll 648–660 Coral 571 Coumaphos 571 fluorescent whitening agents 629, 630 fulvic acid 639–647 humic acid 639–647 mineral oil/petroleum spills 194–196, 214–218 nitriloacetic acid 466 organoboron compounds 807, 808 organotin compounds 761, 762, 767, 768 phenols 277 plant pigments 648–660 polyaromatic compounds 155–157, 166–170, 175 thiabendazole 606, 607 Fluorescent whitening agents, determination of 629, 630 Formaldehyde, determination of 234 Formic acid, determination of 234 Fulvic acid, determination of 639–647 Gas chromatography, general 34–43 Gas chromatography of Abate 569, 570 alcohols 249, 250 Aldrin 517 aliphatic amines 434–436 aliphatic halogen compounds 349, 350, 355– 358 aliphatic hydrocarbons 110, 111, 126–129, 136, 137 alkyl phosphates 477 amides 444–446 amino acids 453 aminophenols 440 anionic detergents 303, 304 aromatic amines 437–439 aromatic hydrocarbons 141, 144 carbamate insecticides/ herbicides 576–580, 582 carbohydrates 258, 259, 261 carboxylic acids 234, 235, 241–247 chlorinated insecticides 496–510,
513, 519–532, 534–540 chloroaromatic amines 440 4(chloromethylsulphenyl)bromobenzene 604 chlorophenols 407–409, 412–416 coprostanol 628, 629 Dalapon 607, 608 DDT 516, 517 Dicamba 601 Dichlorvos 515, 516 Dieldrin 518 diethanolamine 470 dimethyl sulphoxide 489–492 dioxins 422 Diquat 604, 605 disulphides 484–487 Dursban 568, 569 Endosulphan 517, 518 Endrin 518 esters 253–254, 257 ethylnyloestradiol 630 Fenitrothion 570, 571 geosmin 632 glycols 250 haloaromatic compounds 394 halocarboxylic acids 417 haloforms 358, 359, 367–383, 393, 394 hexachlorobenzene 516 hexachlorocyclohexane 516 hexachlorophane 517 2-isobutyl-3methoxypyrazine 632, 633 ketones 253 Malathion 568 mercaptans 484– 487 mestranol 630 methoxychlor 518, 519 2-methylisoborneol 632 Nmethyl pyrrolidone 470 mineral oil/petroleum spills 183–189, 201– 210, 217 multiorganic mixtures 644– 667, 698, 699, 701, 702, 708, 709 nitriles 444 nitrilacetic acid 462, 463, 465, 466, 468, 469 nitrocompounds 441 nitrophenols 443 nitrosamines 448, 451, 452 non-ionic detergents 330–332 organoarsenic compounds 722, 723 organolead compounds 729–731 organomercury compounds 735–737, 746, 747, 755 organophosphorus compounds 543–560, 572–574, 801 organoselenium 771 organosilicon compounds 770 organotin compounds 756–759 ozonization products 634–638 Paraquat 604, 605 Parathion 567, 568 pesticide/ herbicide mixtures 608, 609 phenols 268–273, 280, 283, 285 phenoxyacetic acid herbicides 595– 597, 601 Picloram 602 polyaromatic hydrocarbons 146–152, 159–161, 174–176 polybromophenyls 422 polychlorobiphenyls 396–399, 403– 406 pyrethrin 574, 575 pyridazinones 470 pyridine 470
Index
saturated aliphatic chloro compounds 339–342, 347–349 Squoxin 628 tetrahydrothiophen 494 Toxaphene 519 total halogen 785, 786 triazine herbicides 590, 591, 593, 594 Trichlorphon 517 urea herbicides 584, 585 Gas chromatography–mass spectrometry, general 43–52 Gas chromatography–mass spectrometry of aliphatic amines 435, 436 5-alkylhydantoin 470 benzothiazole 488, 489 chlorinated insecticides 510, 540–542 chloroamino acids 455 chloroanisoles 422 chloromethyl ether 422 4(chloromethylsulphenyl)bromobenzene 604 chlorophenols 415, 417 dialkylhydantoin 470 dioxins 421 geosmin 631–633 2mercaptobenzothiazole 488, 489 2(4-morpholinyl)benzothiazole 488, 489 multiorganic mixtures 664, 683, 697, 699–706, 709, 710 nitrocompounds 441, 442 nitrosamines 451 organolead compounds 733, 734 organotin compounds 759–761 pesticide/ herbicide mixtures 609, 610 phenols 273 polychlorobiphenyls 399, 400, 406 polychloroterphenyls 422, 423 total halogens 785 triazine herbicides 591, 592 Gas stripping methods, general 54 Gas stripping methods, determination of aliphatic hydrocarbons 111, 112, 127, 137 multiorganic mixtures 695 Gel permeation chromatography, determination of calcium lignosulphonate 493 carbohydrates 261 fulvic acid 639–647 haloforms 364 humic acid 639–647 mineral oil/petroleum spills 210 multiorganic mixtures 711 Geosmin, determination of 631–633 Glyphosate, determination of 603 Greases, determination of 219 Growth regulators, determination of 607, 608 Guanazoles, determination of 470
879
Haloaromatic compounds, determination of 394, 395 Halocarboxylic acids, determination of 417–419 Haloforms determination of 358–394 preconcentration of 364, 365, 393 Headspace analysis, general 52–54 Headspace analysis, determination of aliphatic hydrocarbons 111, 112, 126, 127 haloforms 359–361, 384– 387, 393 multiorganic mixtures 695, 701, 702 saturated aliphatic halogen compounds 342, 343 unsaturated aliphatic halogen compounds 351 Herbicides, mixtures of, determination 608–613 Hexachlorobenzene, determination of 516 Hexachlorocyclohexane, determination of 516 Hexachlorophane, determination of 517 High-performance liquid chromatography, general 56–65 High-performance liquid chromatography of aliphatic hydrocarbons 115, 116 amides 445– 447 amino acids 453–455 amino phenols 440 anionic detergents 314 aromatic amines 437, 438 benzothiazole 488, 489 carbamate insecticides/herbicides 581–583 carboxylic acids 236–238, 241, 242 chlorinated insecticides 512 chlorinated polyaromatics 395 chlorophenols 410, 417 chlorophyll 648–660 cobalamin 638, 639 dioxins 421–422 esters 254–257 ethers 289 ethylenediamine tetraacetic acid 461 haloforms 364, 392 2mercaptobenzothiazole 488, 489 monoalkyl quaternary compounds 470 2-(4-morpholinyl)benzothiazole 488, 489 multiorganic mixtures 667– 672, 697, 701, 706, 710–712 nitro compounds 440, 442 non-ionic detergents 325, 334 organic oxidants 633 organocopper compounds 770 organolead compounds 731 organotin compounds 761, 765, 766 ozonization products 634–636
880
Index
Paraquat 604, 605 Parathion 567, 568 pesticide/herbicide mixtures 611, 612 phenols 274, 283–285 phenoxyacetic acid herbicides 599 plant pigments 648–660 polychlorobiphenyls 401, 403 Pyrazon 602, 603 saturated aliphatic chloro compounds 344, 345 thiafluoron 493 unsaturated aliphatic hydrocarbons 354 High-performance liquid chromatography–mass spectrometry, general 65, 66 High-performance liquid chromatography–mass spectrometry of chlorophenols 415 diazo compounds 470 pesticide/ herbicide mixtures 612 Humic acid determination of 639–647 preconcentration of 645, 646 Hydroxymethane sulphinite salts 487, 488 Inductively coupled plasma atomic emission spectrometry, general 17– 23 Inductively coupled plasma atomic emission spectrometry of organoboron compounds 806 organomercury compounds 741, 750 organophosphorus compounds 803 organosilicon compounds 771 Infrared spectroscopy, general 12–13 Infrared spectroscopy of aliphatic hydrocarbons 116–123, 129–132, 139 anionic detergents 304, 312–314 aromatic hydrocarbons 142 Smethyl Fenitrothion 570, 571 1methyl-2-pyrrolidone 470 mineral oil/petroleum spills 193–194, 211– 214 multiorganic mixtures 672, 673, 706 total organic carbon 825, 826 Inositol phosphate, determination of 481 Iodine, determination of 784, 785 Ion chromatography, general 67–70 Ion chromatography of bromine 783 carboxylic acids 238–240 chlorine 778, 779 chlorophenols 411 organic sulphur 805 total organic carbon 819
Ion exchange chromatography, general 66, 67 Ion exchange chromatography of organoarsenic compounds 723 organoboron compounds 807 multiorganic mixtures 711 nonionic detergents 325–327 saturated aliphatic chloro compounds 345, 347 Ion exclusion chromatography of carboxylic acids 238, 241 ozonization products 633–638 Ion pair chromatography of multiorganic mixtures 697 2-Isobutyl–3-methoxypyrazine, determination of 632, 633 Isopropylmethylphosphonofluoridate, determination of 631 Isotachoelectrophoresis, general 67 Isotachoelectrophoresis, determination of Chloromequat 608 Isotope dilution methods, general 33 Isotope dilution methods, determination of organic nitrogen 790–793 Isotope exchange in organic phosphorus 802 Ketones, determination of 253 Kjeldahl digestion, organic nitrogen 786, 795–797 Lactams, determination of 261, 262 Lactic acid, determination of 232–234 Luminescence, general 7–11 Luminescence, determination of adenosine triphosphate 479, 480 2-Lutidine, determination of 470 Malathion, determination of 568 Malic acid, determination of 233 Mandelic acid, determination of 233 Mass spectrometry, general 43–52, 65, 66 Mass spectrometry of geosmin 631–637 halocarboxylic acids 418 S-methyl Fenitrothion 570, 571 multiorganic mixtures 690 non-ionic detergents 332 organophosphorus insecticides 573 organosilicon compounds 770, 771 pesticide/herbicide mixtures 612 triazine herbicides 593
Index
unsaturated aliphatic chloro compounds 350, 355 Mercaptans, determination of 484, 489 2-Mercaptobenzothiazole, determination of 488, 489 Mestranol, determination of 630 Metals in oils, determination of 197– 198, 216 Methacrylic acid, determination of 233, 234 Methoxychlor, determination of 518, 519 S-Methyl Fenitrothion, determination of 570, 571 2-Methylisoborneol, determination of 632 N-Methyl pyrrolidone, determination of 470 Microbial electrodes 863, 870 Mineral oil/petroleum spills analysis of 178–218 preconcentration of 201 Monoalkyl quaternary compounds, determination of 470 2-(4-Morpholinyl)benzothiazole, determination of 488 Multiorganic mixtures analysis of 644– 712 preconcentration of 677–683, 697, 698, 701, 702, 708, 711, 712 Neutron activation analysis, general 24 Neutron activation analysis of bromine 783, 784 chlorine 779, 780 iodine 784, 785 metals in oils 197, 198 organomercury compounds 741 Nitriles, determination of 444 Nitriloacetic acid, determination of 462–471 Nitro compounds determination of 440–442 preconcentration of 442 Nitrophenols, determination of 443, 444 Nitrosamines, determination of 447– 453 Non-ionic detergents determination of 318–334 preconcentration of 330, 331 Nuclear magnetic resonance spectroscopy, general 24 Nuclear magnetic resonance spectroscopy of chlorinated insecticides 512 multiorganic mixtures 675, 701 nitrosamines 448
881
Nucleic acid, determination of 455–457 Organic nitrogen, determination of 786–797, 804, 805 Organic oxidants, determination of 633 Organoantimony compounds, determination of 769 Organoarsenic compounds determination of 772–779 preconcentration of 728 Organoboron compounds, determination of 806, 807 Organocadmium compounds, determination of 770 Organocopper compounds, determination of 770 Organogermanium compounds, determination of 769 Organolead compounds determination of 729–734 preconcentration of 731 Organomercury compounds determination of 733–755 preconcentration of 744, 745, 751 Organophosphorus compounds chemiresistant sensor 482 determination of 482, 542–574, 797– 805 preconcentration of 566 supported Cu-CuO island film 482 surface acoustic wave detector 482 Organoselenium compounds, determination of 771 Organosilicon compounds, determination of 770, 771 Organosulphur compounds determination of 494, 805, 806 surface acoustic wave detector 494 Organotin compounds determination of 755–769 preconcentration of 769 Oscillography, determination of metals in oils 198 Ozonization products, determination of 633–638 Paper chromatography of aliphatic hydrocarbons 125 mineral oil/ petroleum spills 210 Paper electrophoresis of phenoxyacetic acid herbicides 598, 599 Paraquat, determination of 604, 605 Parathion, determination of 567, 568 Particulate organic carbon, determination of 830, 831
882
Index
Pesticide/herbicide mixtures, preconcentration of 612, 613 Pesticide mixtures, determination of 608–613 Phenolic acids, determination of 288 Phenols determination of 263–288 preconcentration of 278, 279 Phenoxyacetic acid herbicides, determination of 594–601 Phenylglyoxylic acid, determination of 233 Photoacoustic spectrometry of organomercury compounds 741 Picloram, determination of 602 Picoline, determination of 470 Plant pigments, determination of 648– 660 Polarography, general 2–5 Polarography of aldehydes 251 amides 445 anionic detergents 305, 307, 311, 314, 315 carboxylic acids 240 chlorophenols 411 esters 257 ethylenediamine tetraacetic acid 460 fulvic acid 639–647 glyphosate 603 humic acid 639–647 hydromethanesulphinite salts 487, 488 lactams 262 nitriloacetic acid 465–467, 469 nitrophenols 444 nonionic detergents 327 organoantimony compounds 769 organoarsenic compounds 723 organolead compounds 731 organophosphorus insecticides 564 Paraquat 604, 605 polyaromatic hydrocarbons 170 polychlorobiphenyls 401, 402 quinones 263 Polyaromatic hydrocarbons determination of 145–178 preconcentration of 155–157, 170– 174, 177, 178 Polybromobiphenyls, determination of 422 Polychlorobiphenyls determination of 395–406, 524–542 preconcentration of 402, 403 Polychloroterphenyls, determination of 422–123 Potentiometric method, determination of anionic detergents 305 carboxylic acids 240 Preconcentration, general 77–86
Preconcentration of aldehydes 251, 252 aliphatic hydrocarbons 129, 136 amino acids 455 anionic detergents 312, 314 aromatic amines 438 aromatic hydrocarbons 144 carbamate insecticides/herbicides 582 carboxylic acids 241 chlorinated insecticides 514, 515, 522, 533–534 chlorophenols 412, 417 fulvic acids 645, 646 haloforms 364, 365, 393 humic acids 645, 646 mineral oil/ petroleum spills 201 multiorganic mixtures 677–683, 697, 698, 701, 702, 708, 711, 712 nitro compounds 442 non-ionic detergents 330, 331 organoarsenic compounds 728 organolead compounds 732 organomercury compounds 744, 745, 751 organotin compounds 769 pesticide/herbicide mixtures 612, 613 phenols 278 phosphorus insecticides 566 polyaromatic hydrocarbons 155–157, 170–174, 177–178 polychlorobiphenyls 403, 403 saturated aliphatic chloro compounds 346, 347 urea herbicides 589 Purge and trap analysis, general 54 Purge and trap analysis, determination of aldehydes 252 haloforms 362, 363, 387–391 saturated aliphatic halogen compounds 343, 344 unsaturated aliphatic halogen compounds 351–355 Pyrazone, determination of 602, 603 Pyrethrin insecticides, determination of 574 Pyridazinones, determination of 470 Pyridine, determination of 470 Pyrolysis–gas chromatography, general 54-56 Pyrolysis–gas chromatography of chlorolignosulphonic acids 420, 493, 494 Paraquat 604, 605 total organic carbon 816, 817 Quinones, determination of 263 Raman spectroscopy, general 12, 13 Raman spectroscopy of mineral oil/ petroleum spills 211, 214 multiorganic mixtures 673 phenols
Index
275–277, 281 phosphorus insecticides 564 Reverse osmosis of multiorganic mixtures 689, 690 Saturated aliphatic halogen compounds determination of 339– 349 preconcentration of 346, 347 Spectrophotometric methods, general 5–7 Spectrophotometric determination of alcohols 248, 249 aldehydes 250, 252, 253 aliphatic amines 434–436 aliphatic hydrocarbons 135 amides 446 amino triazole 470 anionic detergents 299–301, 307, 310, 312, 314 aromatic amines 437–439 aromatic hydrocarbons 141 benethonium salts 487 carbamate insecticides/herbicides 575, 576 carbohydrates 258, 261 carboxylic acids 232–234, 243–245 carboxymethyl succinate 630, 631 cationic detergents 315, 316 chemical oxygen demand 840, 843, 844 chlorinated isocyanurates 420 chlorophenols 407 chlorophyll 648– 660 2,3-dichloro-1,4naphthaquinone 631 dioxans 262 glycols 250 guanazole 470 hydrazine 470 iodine 784 nitriles 444 nitriloacetic acid 462 nitro compounds 440–442 non-ionic detergents 318–320, 331, 332, 334 nucleic acid 455, 458 organic nitrogen 187, 793–795 organoarsenic compounds 723, 727, 728, 729 organoboron compounds 807, 808 organolead compounds 733 organophosphorus compounds 799–801, 803, 804 organotin compounds 755, 756 Paraquat 604, 605 Parathion 567, 568 phenols 264– 268, 279, 280–285 phosphorus insecticides 542, 543, 571, 572 plant pigments 648–660 Pyrazon 602, 603 Squoxin 628 Thiram 575 Toxaphene 519 Squoxin, determination of 628 Substituted urea herbicides, determination of 583–589
883
Supercritical fluid extraction, general 71–76 Supercritical fluid extraction, determination of chlorinated insecticides 532 multiorganic mixtures 673–675 polychlorobiphenyls 532 Supported Cu–CuO island film detector 482 Surface acoustic wave detector, determination of organophosphorus compounds 482 organosulphur compounds 494 Tetrahydrothiophen, determination of 494 Thiabendazole, determination of 606, 607 Thiafluoron, determination of 493 Thin-layer chromatography, general 75 Thin-layer chromatography of Abate 569, 570 aldehydes 251 aliphatic hydrocarbons 123–125, 132–134, 137, 138 aromatic hydrocarbons 142–144 carbamate insecticides/ herbicides 580, 581 carboxylic acids 235 chlorinated insecticides 510– 512, 523, 524 chlorophyll 410, 648– 660 ethynyloestradiol 630 lactams 261 Malathion 568 mestranol 630 mineral oil/petroleum spills 189– 193 multiorganic mixtures 706 nitrophenols 443 non-ionic detergents 322 organomercury compounds 754, 755 organotin compounds 761 pesticide/herbicide mixtures 610 phenolic acids 288 phenols 273, 274, 280, 282 phenoxyacetic acid herbicides 598 phosphorus insecticides 560–562, 574 plant pigments 648–660 polyaromatic compounds 155, 161 polychlorobiphenyls 400, 401 Pyrazon 602, 603 quinones 263 saturated aliphatic chloro compounds 343 Thiram insecticides, determination of 575 triazine herbicides 592 urea herbicides 585, 586 Titration, general 1–2 Titration of aliphatic amines 436 amides 446 anionic detergents 298,
884
Index
299 bromine 784 cationic detergents 315, 317, 318 chemical oxygen demand 838–840, 842–844 chlorine 778, 781–783 disulphides 484 fulvic acids 639–647 humic acids 639–647 iodine 785 mercaptans 484 nitriles 444 nitriloacetic acid 467, 468 nucleic acid 458 total nitrogen 795, 796 total oxygen demand 853 Total halogen, determination of 785, 786 Total inorganic carbon, determination of 824, 826 Total organic carbon, determination of 808–824 Total oxygen demand, determination of 850–854 Toxaphene, determination of 519 Triazine herbicides, determination of 589–594 Trichlorophon, determination of 517 Ultraviolet digestion, general 7 Ultraviolet digestion, determination of dissolved organic carbon 827 total organic carbon 814, 815, 819 Ultraviolet irradiation, determination of organic nitrogen 787–790 Ultraviolet spectroscopy,, general 7 Ultraviolet spectroscopy, determination of aliphatic hydrocarbons 139, 140 anionic
detergents 304 aromatic hydrocarbons 142 calcium lignosulphonate 493 cationic detergents 316 fulvic acid 639–647 humic acid 639–647 2-lutidene 470 S-methyl Fenitrothion 570, 571 mineral oil/petroleum spills 214 phenols 277, 284 picolene 470 saturated aliphatic chloro compounds 345 unsaturated aliphatic halogen compounds 349– 358 Urea herbicides determination of 470 preconcentration of 589 Voltammetry, determination of chlorinated insecticides 532 polychlorobiphenyls 532 Thiram 575 Wet oxidation, determination of dissolved organic carbon 826, 827, 829–830 total organic carbon 813, 814, 818, 819 X-ray fluorescence spectroscopy,,general 25–33 X-ray fluorescence spectroscopy, determination of alkyl phosphates 478 aryl phosphates 478 metals in oils 198
