Handbook of Water Analysis

HANDBOOK OF

WATER ANALYSIS SECOND EDITION

HANDBOOK OF

WATER ANALYSIS SECOND EDITION

EDITED BY

LEO M. L. NOLLET

Boca Raton London New York

CRC Press is an imprint of the Taylor & Francis Group, an informa business

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2007 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-10: 0-8493-7033-7 (Hardcover) International Standard Book Number-13: 978-0-8493-7033-5 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http:// www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data Handbook of water analysis / editor, Leo M.L. Nollet. -- 2nd ed. p. cm. Includes bibliographical references and index. ISBN-13: 978-0-8493-7033-5 (alk. paper) ISBN-10: 0-8493-7033-7 (alk. paper) 1. Water--Analysis--Handbooks, manuals, etc. I. Nollet, Leo M. L., 1948- II. Title. QD142.H36 2007 628.1’61--dc22 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

2007006719

Table of Contents

Preface ........................................................................................................................................... vii Author ............................................................................................................................................ ix Contributors .................................................................................................................................. xi 1.

Sampling Methods in Surface Waters .............................................................................1 Munro Mortimer, Jochen F. Mu¨ller, and Matthias Liess

2.

Methods of Treatment of Data ........................................................................................47 Riccardo Leardi

3.

Radioanalytical Methodology for Water Analysis ......................................................77 Jorge S. Alvarado

4.

Bacteriological Analysis of Water ...................................................................................97 Paulinus Chigbu and Dmitri Sobolev

5.

Marine Toxins Analysis ..................................................................................................135 Luis M. Botana, Amparo Alfonso, M. Carmen Louzao, Mercedes R. Vieytes, and Marı´a R. Velasco

6.

Halogens .............................................................................................................................157 Geza Nagy and Livia Nagy

7.

Analysis of Sulfur Compounds in Water....................................................................201 Laura Coll and Leo M.L. Nollet

8.

Phosphates..........................................................................................................................219 Philippe Monbet and Ian D. McKelvie

9.

Cyanides .............................................................................................................................253 Meissam Noroozifar

10.

Asbestos in Water .............................................................................................................269 James S. Webber

11.

Heavy Metals, Major Metals, Trace Elements............................................................275 Jorge E. Marcovecchio, Sandra E. Botte´, and Rube´n H. Freije

12.

Determination of Silicon and Silicates ........................................................................313 Salah M. Sultan v

vi 13.

Main Parameters and Assays Involved with Organic Pollution of Water...........337 Claudia E. Domini, Lorena Vidal, and Antonio Canals

14.

Determination of Organic Nitrogen and Urea ...........................................................367 Stefano Cozzi and Michele Giani

15.

Organic Acids ....................................................................................................................393 Sigrid Peldszus

16.

Determination of Phenolic Compounds in Water.....................................................409 ˚ ke Jo¨nsson Tarekegn Berhanu and Jan A

17.

Characterization of Freshwater Humic Matter ...........................................................435 Juhani Peuravuori and Kalevi Pihlaja

18.

Analysis of Pesticides in Water .....................................................................................449 Evaristo Ballesteros Tribaldo

19.

Fungicide and Herbicide Residues in Water..............................................................491 Sara Bogialli and Antonio Di Corcia

20.

Polychlorobiphenyls ........................................................................................................529 Alessio Ceccarini and Stefania Giannarelli

21.

Determination of PCDDs and PCDFs in Water.........................................................563 Luigi Turrio-Baldassarri, Anna L. Iamiceli, and Silvia Alivernini

22.

Polynuclear Aromatic Hydrocarbons ...........................................................................579 Chimezie Anyakora

23.

Analysis of Volatile Organic Compounds in Water .................................................599 Iva´n P. Roma´n Falco´ and Marta Nogueroles Moya

24.

Analysis of Surfactants in Samples from the Aquatic Environment ....................667 B. Thiele and Leo M.L. Nollet

25.

Analysis of Endocrine Disrupting Chemicals and Pharmaceuticals and Personal Care Products in Water...........................................................................693 Guang-Guo Ying

26.

Residues of Plastics..........................................................................................................729 Caroline Sablayrolles, Mireille Montre´jaud-Vignoles, Michel Treilhou, and Leo M.L. Nollet

Index .............................................................................................................................................745

Preface

The Handbook of Water Analysis, Second Edition, discusses as in the first edition, all types of water: freshwater from rivers, lakes, canals, and seawater, as well as groundwater from springs, ditches, drains, and brooks. Most of the chapters describe the physical, chemical, and other relevant properties of water components, and covers sampling, cleanup, extraction, and derivatization procedures. Older techniques that are still frequently used are compared to recently developed techniques. The reader is also directed to future trends. A similar strategy is followed for discussion of detection methods. In addition, some applications of analysis of water types (potable water, tap water, wastewater, seawater) are reviewed. Information is summarized in graphs, tables, examples, and references. Because water is an excellent solvent, it dissolves many substances. To get correct results and values, analysts have to follow sample strategies. Sampling has become a quality-determining step (Chapter 1). Statistical treatment of data ensures the reliability of the results. Statistical and chemometrical methods are discussed in Chapter 2. Chapter 3 discusses new technologies on radionuclides and their possible health hazards in water and the whole environment. Water is a living element, housing many organisms—wanted or unwanted, harmful or harmless. Some of these organisms produce toxic substances. Chapter 4 and Chapter 5 discuss bacteriological and algal analysis. Humans consume and pollute large quantities of water. Chapter 6 through Chapter 26 cover injurious or toxic substances of domestic, agricultural, and industrial sources: halogens, sulphur compounds, phosphates, cyanides, asbestos, heavy and other metals, silicon compounds, nitrogen compounds, organic acids, phenolic substances, humic matter, pesticides, insecticides, herbicides, fungicides, PCBs, PCDFs, PCDDs, PAHs, VOCs, surfactants, EDCs, and plastics residues. Chapter 23, Chapter 25, and Chapter 26 discuss in detail the separation and analysis of volatile organic compounds (VOCs), endocrine disrupting compounds (EDCs) and pharmaceutical and personal care products (PPCPs), and plastics residues, respectively. Many of these compounds are widely distributed in the environment but in very small quantities. This book may be used as a primary textbook for undergraduate students learning techniques of water analysis. Furthermore, it is intended for the use of graduate students involved in the analysis of water. All contributors are international experts in their field of water analysis. I would like to thank them cordially for all their efforts. This work is dedicated to my three granddaughters: Fara, Fleur, and Kato. I hope they will live on a blue planet, the blue being the color of healthy water. Leo M.L. Nollet

vii

Author

Leo M.L. Nollet is a professor of biochemistry, aquatic ecology, and ecotoxicology in the department of applied engineering sciences, University College Ghent, member of Ghent University Association, Ghent, Belgium. His main research interests are in the areas of food analysis, chromatography, and analysis of environmental parameters. He is author or coauthor of numerous articles, abstracts, and presentations, and is the editor of Handbook of Food Analysis, 2nd ed. (three volumes), Food Analysis by HPLC, 2nd ed., Handbook of Water Analysis (all titles, Marcel Dekker, Inc.), Chromatographic Analysis of the Environment, 3d ed., Advanced Technologies of Meat Processing, and Radionuclide Concentrations in Food and the Environment (all titles, CRC Press, Taylor & Francis). He received his MS (1973) and PhD (1978) in biology from the Katholieke Universiteit Leuven, Leuven, Belgium.

ix

Contributors

Amparo Alfonso Departamento de Farmacologı´a, Universidad de Santiago de Compostela, Lugo, Spain Silvia Alivernini Dipartimento di Sanita` Alimentare ed Animale, Istituto Superiore di Sanita`, Rome, Italy Jorge S. Alvarado Environmental Science Division, Argonne National Laboratory, Argonne, Illinois Chimezie Anyakora Department of Pharmaceutical Chemistry, University of Lagos, Lagos, Nigeria Tarekegn Berhanu Department of Chemistry, University of Addis Ababa, Addis Ababa, Ethiopia Sara Bogialli Dipartimento di Chimica, Universita ‘‘La Sapienza’’, Rome, Italy Luis M. Botana Departamento de Farmacologı´a, Universidad de Santiago de Compostela, Lugo, Spain Sandra E. Botte´ Area de Oceanografı´a Quı´mica, Instituto Argentino de Oceanografı´a – CONICET, Bahı´a Blanca, Argentina Antonio Canals Departamento Quı´mica Analı´tica, Nutricio´n y Bromatologia, Universidad de Alicante, Alicante, Spain Alessio Ceccarini Department of Chemistry and Industrial Chemistry, Universita degli Studi di Pisa, Pisa, Italy Paulinus Chigbu Department of Natural Sciences, University of Maryland Eastern Shore, Princess Anne, Maryland Laura Coll Departamento de Ingenierı´a Quı´mica, University of Valencia, Burjassot, Spain Antonio Di Corcia Dipartimento di Chimica, Universita degli Studi di Roma ‘‘La Sapienza’’ Piazzale Aldo Moro, Rome, Italy Stefano Cozzi Consiglio Nazionale delle Ricerche, Istituto di Scienze Marine, Sede di Trieste, Trieste, Italy

xi

xii

Claudia E. Domini Departamento de Quı´mica, Universidad Nacional del Sur, Bahı´a Blanca, Argentina Iva´n P. Roma´n Falco´ Departamento de Quı´mica Analı´tica Nutricio´n y Bromatologia, Universidad de Alicante, Alicante, Spain Rube´n H. Freije Departamento de Quı´mica, Universidad Nacional del Sur, Bahı´a Blanca, Argentina Michele Giani Laboratory of Marine Biogeochemistry and Chemical Oceanography, Istituto Centrale per la Ricerca scientifica e tecnologica Applicata al Mare, Chioggia (Venice), Italy Stefania Giannarelli Department of Chemistry and Industrial Chemistry, Universita degli Studi di Pisa, Pisa, Italy Anna L. Iamiceli Dipartimento di Sanita` Alimentare ed Animale, Istituto Superiore di Sanita`, Rome, Italy ˚ ke Jo¨nsson Jan A Sweden

Department of Analytical Chemistry, Lund University, Lund,

Riccardo Leardi Department of Chemistry and Food and Pharmaceutical Technologies, University of Genoa, Genoa, Italy Matthias Liess Department of System Ecotoxicology, VFZ-Helmholtz Centre for Environmental Research Permoserstrasse 15, D-04318 Leipzig, Germany M. Carmen Louzao Departamento de Farmacologı´a, Universidad de Santiago de Compostela, Lugo, Spain Jorge E. Marcovecchio Area de Oceanografı´a Quı´mica, Instituto Argentino de Oceanografı´a – CONICET, Bahı´a Blanca, Argentina Ian D. McKelvie Water Studies Centre, School of Chemistry, Monash University, Victoria, Australia Philippe Monbet Water Studies Centre, School of Chemistry, Monash University, Victoria, Australia Mireille Montre´jaud-Vignoles Laboratoire Chimie Agro-Industrielle, UMR 1010 INRA/INP-ENSIACET, TOULOUSE, France Munro Mortimer Environmental Protection Agency, Brisbane, Australia Marta Nogueroles Moya Departamento de Quı´mica Analı´tica Nutricio´n y Bromatologia, Universidad de Alicante, Alicante, Spain

xiii

Jochen F. Mu¨ller National Research Centre for Environmental Toxicology, The University of Queensland, Brisbane, Australia Geza Nagy Department of General and Physical Chemistry, University of Pecs, Pecs, Hungary Livia Nagy Department of General and Physical Chemistry, University of Pecs, Pecs, Hungary Leo M.L. Nollet Department of Engineering Sciences, Hogeschool Gent, Ghent, Belgium Meissam Noroozifar Analytical Research Laboratory, Department of Chemistry, Faculty of Science, University of Sistan and Baluchestan (USB), Zahedan, Iran Sigrid Peldszus Department of Civil and Environmental Engineering, University of Waterloo, Waterloo, Ontario, Canada Juhani Peuravuori Department of Chemistry, Laboratory of Organic Chemistry and Chemical Biology, University of Turku, Turku, Finland Kalevi Pihlaja Department of Chemistry, Laboratory of Organic Chemistry and Chemical Biology, University of Turku, Turku, Finland Caroline Sablayrolles Laboratoire Chimie Agro-Industrielle, UMR 1010 INRA/ INP-ENSIACET, TOULOUSE, France Dmitri Sobolev Department of Biology, Jackson State University, Jackson, Mississippi Salah M. Sultan Department of Chemistry, King Fahd University, Dhahran, Saudi Arabia B. Thiele Institute for Chemistry and Dynamics of the Geosphere, Institute III: Phytosphere, Research Centre Ju¨lich, Ju¨lich, Germany Michel Treilhou Laboratoire de Chimie et Biochimie des Interactions, Centre Universitaire Jean-Franc¸ois Champollion, Albi, France Evaristo Ballesteros Tribaldo Department of Physical and Analytical Chemistry, E.P.S. of Linares, University of Jae´n, Jae´n, Spain Luigi Turrio-Baldassarri Dipartimento di Sanita` Alimentare ed Animale, Istituto Superiore di Sanita`, Rome, Italy

xiv

Marı´a R. Velasco, EU-Community Reference Laboratory on Marine Biotoxins, Agencia Espan˜ola de Seguridad Alimentaria, Vigo, Spain Lorena Vidal Departamento Quı´mica Analı´tica Nutricio´n y Bromatologia, Universidad de Alicante, Alicante, Spain Mercedes R. Vieytes Departamento de Farmacologı´a, Universidad de Santiago de Compostela, Lugo, Spain James S. Webber Wadsworth Center, New York State Department of Health, Albany, New York Guang-Guo Ying CSIRO Land and Water, Adelaide Laboratory PMB 2, Glen Osmond, Australia and, Guangzhou Institute of Geochemistry, Chinese Academy of Sciences, Guangzhou, China

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1 Sampling Methods in Surface Waters Munro Mortimer, Jochen F. Mu¨ller, and Matthias Liess

CONTENTS 1.1 Introduction ........................................................................................................................... 2 1.2 General Aspects of Sampling and Sample Handling ..................................................... 3 1.2.1 Initial Considerations ............................................................................................. 3 1.2.2 Spatial Aspects ........................................................................................................ 3 1.2.3 Temporal Aspects ................................................................................................... 3 1.2.4 Number of Samples................................................................................................ 5 1.2.5 Sample Volume ....................................................................................................... 5 1.2.6 Storage and Conservation ..................................................................................... 6 1.2.6.1 Contamination.......................................................................................... 6 1.2.6.2 Loss ............................................................................................................ 6 1.2.6.3 Sorption ..................................................................................................... 7 1.2.6.4 Recommended Storage ........................................................................... 8 1.2.6.5 Quality Control in Water Sampling...................................................... 8 1.3 Sampling Strategies for Different Ecosystems ................................................................. 8 1.3.1 Lakes and Reservoirs ........................................................................................... 13 1.3.2 Streams and Rivers ............................................................................................... 15 1.3.2.1 Location of Sampling within the Stream ........................................... 15 1.3.2.2 Description of the Longitudinal Gradient ......................................... 15 1.3.2.3 Temporal Changes of Water Quality ................................................. 16 1.3.2.4 Using Sediments to Integrate over Time ........................................... 17 1.3.3 Estuarine and Marine Environments ................................................................ 17 1.3.4 Urban Areas........................................................................................................... 18 1.4 Sampling Equipment.......................................................................................................... 20 1.4.1 General Comments ............................................................................................... 20 1.4.2 Manual Sampling Systems .................................................................................. 20 1.4.2.1 Simple Sampler for Shallow Water..................................................... 20 1.4.2.2 Sampler for Large Quantities in Shallow Water .............................. 20 1.4.2.3 Simple Sampler for Deepwater ........................................................... 20 1.4.2.4 Deepwater Sampler (Not Adding Air to the Sample)..................... 21 1.4.2.5 Deepwater Sampler for Trace Elements (Allowing Air to Mix with the Sample) ....................................................................... 21 1.4.3 Systems for Sampling the Benthic Boundary Layer at Different Depths................................................................................................ 23 1.4.3.1 Deepwater (>50 m) ............................................................................... 23 1.4.3.2 Shallow Water ( 12

K2Cr2O7

Add potassium dichromate 0.05% by mass final concentration

Source: From ISO. Water quality—Sampling—Part 3: Guidance on the preservation and handling of samples. ISO 5667/3 2003.

can be described in relation to either the goals of the study or the ecosystems involved. In the following section, the different sampling strategies appropriate to the main types of ecosystem (still water, flowing water, estuarine or marine environment) and their temporal and spatial scaling are discussed. In Section 1.3.4, considerations for sampling storm water runoff are used as an example of a sampling design specific to urban areas. 1.3.1

Lakes and Reservoirs

Often a number of physical, chemical, and biological processes have to be considered as they may markedly affect water quality and its spatial variations. Sources of heterogeneity within a body of water that need careful consideration in selecting sampling sites are as follows:

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thermal stratification, which leads to variations of quality in depth;

. .

effects of influent streams; lake morphology; and

.

wind.

Any of these factors acting alone or in combination may produce both lateral and vertical heterogeneity of water quality. Some important topics relevant to the choice of sites for sampling are noted (e.g., Refs. [3,18]). It must be kept in mind that shallow and/or relatively isolated embayments of lakes and reservoirs may show marked differences in quality from the main body of water. In water bodies of sufficient depth in temperate climates, thermal stratification is often the most important source of vertical heterogeneity from spring to autumn (Figure 1.3). Measurement of dissolved oxygen and temperature is a convenient means of following the development of such stratification, and has the advantage that both measurements can be made automatically and continuously in situ. Thermal stratification may retard the mixing of streams entering lakes or reservoirs. This important source of materials derived from the surrounding land consequently has to be sampled with due consideration of its spatial variability. The number of algae in the surface layer of a water body may have a marked effect on the concentrations of nutrients and other substances: It is often not possible to measure any dissolved nutrients during algal blooms, because all nutrients are bound in the algae. Therefore, the trophic status and spatial heterogeneity in the distribution of algae should be considered while choosing a sampling site. The choice of the correct sampling point can depend on the depth of a lake [19]. These authors have compared different water sampling techniques in a series of lakes.

Spring

48C O2

Summer

208C O2

E M H 48C

Autumn O2

Winter

Ice FIGURE 1.3 Seasonal variation of oxygen and temperature within the different layers of a meso/eutrophic lake in temperate latitudes (E, epilimnion; M, metalimnion; H, hypolimnion).

O2

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In deep lakes, they observed no significant differences between mean summer nutrient concentrations measured in a tube sample integrating over the photic zone, taken from the deepest point, and a surface dip sample taken by wading into the water’s edge. However, in shallower lakes the integrating tube sampler gave significantly higher estimates of mean concentrations than the other method due to the increase in volume of the unmixed hypolimnion, which reduced the depth of the well-mixed epilimnion to less than the tube length. For national survey purposes they suggested samples taken from the edge of the lake as the most cost effective. 1.3.2

Streams and Rivers

1.3.2.1 Location of Sampling within the Stream Sampling locations especially in larger streams and rivers should, whenever possible, be at cross sections where vertical and lateral mixing of any effluents or tributaries is complete. To avoid nonrepresentative samples caused by surface films and/or the entrainment of bottom deposits, it has often been recommended that samples should, whenever possible, be collected no closer than 30 cm to the surface or the bottom [5]. Simple surface-grab procedures have been compared with more involved, crosssectionally integrated techniques in streams [20]. Paired samples for analysis of selected constituents were collected over various flow conditions at four sites to evaluate differences between the two sampling methods. Concentrations of dissolved constituents were not consistently different. However, concentrations of suspended sediment and the total forms of some sediment-associated constituents, such as phosphorus, iron, and manganese, were significantly lower in the surface-grab samples than in the cross-sectionally integrated samples. The largest median percent difference in concentration for a site was 60% (total recoverable manganese). Median percent differences in concentration for sediment-associated constituents considering all sites grouped were in the range of 20%–25%. The surface-grab samples underrepresented concentrations of suspended sediment and some sediment-associated constituents, thus limiting the applicability of such data for certain purposes. When the quality of river water extracted for a particular end use is of interest (e.g., the production of drinking water), the sampling point should, in general, be at or near the point of extraction. It must be noted, however, that changes in quality may occur between the actual point of extraction and the inlet to the treatment plant. If the amount or time of extraction is to be controlled on the basis of the water quality, an additional sampling location upstream of the extraction point will usually be needed, the distance upstream being dependent on the travel-time of the river, the speed with which the relevant analysis can be made, and the upstream locations of sources of the determinants. This is of course difficult to achieve. The need of an early warning system for drinking water purposes was emphasized by the Sandoz accident in 1986, since which online biomonitors have been in place in the River Rhine [21]. 1.3.2.2 Description of the Longitudinal Gradient When the aim is to assess the quality of a complete stream, river or river basin, the number of potentially relevant sampling locations is usually extremely large. It is, therefore, usually necessary to assign different priorities to the various locations in order to arrive at a feasible sampling program [5,18]. Such considerations are very closely connected to the issue of sampling frequency, and a number of approaches for the overall design of sampling programs for river systems have been described [3]. The value of, and need for, identification of locations where quality problems are or may be most acute,

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have been stressed several times. These questions can be addressed with fixed-location monitoring and intensive, short-term surveys at selected locations for the routine assessment of rivers. Water quality is usually monitored on a regular basis at only a small number of locations in a catchment, generally concentrated at the catchment outlet. This integrates the effect of all the point and nonpoint source processes occurring throughout the catchment. However, effective catchment management requires data which identify major sources and processes. For example, as part of a wider study aimed at providing technical information for the development of integrated catchment management plans for a 5000 km2 catchment in southeastern Australia, a ‘‘snapshot’’ of water quality was undertaken during stable summer flow conditions. These low flow conditions exist for long periods, so water quality at these flow levels is an important constraint on the health of instream biological communities. Over a 4-day period, a study of the low flow water quality characteristics throughout the Latrobe River catchment was undertaken. Sixty-four sites were chosen to enable a longitudinal profile of water quality to be established. All tributary junctions and sites along major tributaries as well as all major industrial inputs were included. Samples were analyzed for a range of parameters including total suspended solid concentration, pH, dissolved oxygen, electrical conductivity, turbidity, flow rate, and water temperature. Filtered and unfiltered samples were taken from 27 sites along the mainstream and tributary confluences for the analysis of total N, NH4 , oxidized N, total P, and dissolved reactive P concentrations. The data were used to illustrate the utility of this sampling methodology for establishing specific sources and estimating nonpoint source loads of phosphorus, total suspended solids, and total dissolved solids. The methodology enabled several new insights into system behavior, including quantification of unknown point discharges, identification of key instream sources of suspended material, and the extent to which biological activity (phytoplankton growth) affects water quality. The costs and benefits of the sampling exercise are reviewed in Ref. [22]. 1.3.2.3

Temporal Changes of Water Quality

The discharge of streams in comparison with larger rivers is highly dynamic, depending mainly on local rainfall conditions and/or groundwater level [23]. It follows that the chemical composition of the stream water is profoundly influenced by the allochthonous input of water, nutrients, sediments, and pesticides. Two-thirds of the contamination of headwater streams with sediments, nutrients, and pesticides is caused by those nonpoint sources [24]. Substances with a high water solubility are introduced through soil filtration. Less water-soluble substances enter by way of the surface water runoff during heavy rains [25]. The total loss of a particular pesticide depends on the time period between application and the rain event, the pattern of precipitation, various soil parameters, and the physical and chemical properties of the pesticide. Consequently, streams with an agricultural catchment area are susceptible to unpredictable, brief pesticide inputs following precipitation [26,27]. To determine the influence of sampling frequency on the reliability of water quality estimates in small streams, a cultivated (0:12 km2 ) and a forested basins (0:07 km2 ) were studied in spring and autumn [28]. During the 2-month spring season and 3-month autumn season 97%–99% of the annual loads of total nitrogen, total phosphorus, and suspended solids was acquired from the cultivated basin and 89%–91% from the forested basin. During the same seasons 99% and 87% of the total annual runoffs were recorded in the cultivated and forested basins. This means that in only 5 months of the year more than 95% of the nutrient and water runoff occurred in the cultivated catchment, and about 90% in the forested catchment. Thus the values of nonpoint loads, normally presented as

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annual means, can give a very misleading impression of the effects of nonpoint loading on watercourses, particularly in the case of relatively small streams. The same author [28] estimated the number of samples needed to calculate the load of various substances by varying the sampling frequency at the two sites. In the cultivated basin the means of concentration data would be within +20% of the mean of the whole data set in spring, if nitrogen and phosphorus samples were taken at least five times and suspended-solid samples at least three times monthly. In the forested basin the corresponding sampling frequencies were twice monthly for nitrogen samples, and four times monthly for phosphorus and suspended solids. In autumn the concentration means in runoff waters would be within +20% of the mean obtained using the whole data set if three samples per month were taken for nitrogen and phosphorus and five samples for suspended solids. In the forested basin the same deviation of the mean would be obtained with 1 nitrogen sample, 5 phosphorus samples, and 16 suspended-solid samples. When intending to measure the peak concentrations of slightly soluble substances in streams within a cultivated watershed, it is necessary to use runoff-triggered sampling methods. A headwater stream in an agricultural catchment in northern Germany was intensively monitored for insecticide occurrence (lindane, parathion-ethyl, fenvalerate). Brief insecticide inputs following precipitation with subsequent surface runoff result in high concentrations in water and suspended matter (e.g., fenvalerate: 6:2 mg L1 , 302 mg kg1 ). These transient insecticide contaminations are typical of headwater streams with an agricultural catchment area, but have been rarely reported. Event-controlled sampling methods for the determination of this runoff-related contamination with a time resolution of as little as 1 h make it possible to detect such events [29]. Within monitoring programs, loading errors are generally associated with an inadequate specification of the temporal variance of discharge and of the parameters of interest. Often little consideration is given to the impact of additional transport characteristics on contaminant sampling error and design. Detailed examination of five transport characteristics at a single river cross section emphasizes the importance of understanding the complete transport/loading regime at a sampling station, defining the required end products of the monitoring program, and defining the accuracy required to meet specific program needs before implementing or evaluating a monitoring program. River transport characteristics are: (a) contaminant transport modes, (b) short-term temporal and seasonal variability, (c) the relationship between dissolved and particulate contaminant concentrations and discharge, (d) load distribution with sediment particle size, and (e) spatial variability in a cross section [30]. It is also worth noting that the procedure known as catchment quality control, though intended for a different purpose, includes the identification of most important effluents entering a river system from a viewpoint of water quality [31]. 1.3.2.4 Using Sediments to Integrate over Time The analysis of river sediments has been suggested as a convenient means of reconnaissance of river systems to decide the locations where water quality is of particular interest with respect to pollutants. 1.3.3

Estuarine and Marine Environments

The potential spatial heterogeneity (lateral and vertical—both are time-dependent) of these bodies of water makes it essential that sampling locations be chosen with reference to the relevant basic processes [18]. Sampling of ocean waters and the handling of such samples have been described in general [4].

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A well-known practical problem is the unintended contamination of samples by material released from the research vessel or by the sampling apparatus. A sampling apparatus for the collection and filtration of up to 28 L of water at sea has been designed to minimize possible contamination from both the equipment and the ship’s surroundings [32]. It was used in the analysis of chlorinated biphenyls (CBs), persistent OC pesticides, and pentachlorophenol (PCP), in both the aqueous and particulate phases. The system is suitable for collection of estuarine and coastal waters where the levels of dissolved CBs, OCs, and PCP are above the limit of determination of 15 pg L1 . The efficiency of the recovery of these compounds and variance of the extraction and analysis have been estimated by analysis of filtered seawater spiked at a range of concentrations from picogram per liter to nanogram per liter. Recoveries ranged from 66.5% to 97.3% with coefficients of variation for the complete method from 7.2% to 29.9%. The procedure of using small boats provided with sample bottles attached to a telescopic device is recommended as a means to minimize contamination from the research vessel during coastal water sampling. Of the wires used to suspend samplers, plasticcoated steel gave negligible, and Kevlar and stainless steel only slight, contamination for some metals [7]. The high spatial and temporal variability of estuaries poses a challenge for characterizing estuarine water quality. This problem was examined by conducting monthly highresolution transects for several water quality variables (chlorophyll-a, suspended particulate matter, and salinity) in San Francisco Bay, California [33]. Using these data, six different ways of choosing station locations along a transect, in order to estimate mean conditions, were compared. In addition, 11 approaches to estimating the variance of the transect mean when stations are equally spaced were compared, and the relationship between variance of the estimated transect mean and number of stations was determined. These results were used to derive guidelines for sampling along the axis of an estuary. In addition, the changes in the concentration of various substances due to the tide seem to be extremely important. Seawater with a low concentration of substances becomes mixed with the highly loaded water in the estuaries and along the shores. Automatic samplers can be used to integrate the concentrations of materials over time (see Section 1.4). An overview of the analysis of polar pesticides in water samples has been presented [34]. The sampling plans and strategies for different types of waters such as rivers, wells, and seawater are discussed. In situ preconcentration methods, involving online techniques or direct measurement, are suggested as alternatives to conventional techniques. Attention is devoted to the influence of organic matter and its interaction with polar pesticides. The use of various types of filtration steps prior to the preconcentration of the analytes from water samples is also reviewed. 1.3.4

Urban Areas

In urban areas, sampling strategies for storm water runoff from industries and municipalities are of specific importance. The United States Federal Storm Water Regulations of 1990 specify protocols for such storm water runoff sampling. These regulations define two separate samples that must be collected when a storm occurs. A first-flush sample is to be collected during the first 30 min of the storm event. A flow-weighted composite sample must be collected for the entire storm event or at least the first 3 h of the event [8]. The first-flush sample and the flow-weighted composite sample must be analyzed for the pollutants listed in Table 1.3. In general, the sample volume required for laboratory analysis depends on the particular pollutants being monitored and varies for each application. As a general rule, a 3 L sample volume for both first-flush and flow-weighted composite sample usually is sufficient for the majority of applications [8].

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TABLE 1.3 Storm Water Analysis Requirements according to the United States Federal Storm Water Regulations Industrial Pollutant

First-Flush Grab Sample

Oil and grease pH Biological oxygen demand, BOD Chemical oxygen demand, COD Total suspended solids, TSS Total phosphorus Nitrate and nitrite nitrogen Total Kjeldahl nitrogen Any pollutant in the facility’s effluent guideline Any pollutant in the facility’s NPDESa permit Any pollutant in the EPA form 2F tables believed to be present Total dissolved solids Fecal streptococcus Fecal coliforms Dissolved phosphorus Total ammonia plus organic nitrogen 13 Metals, total cyanide, and total phenol 28 Volatile compounds 11 Acid compounds 46 Base/neutral compounds 25 Pesticides

Municipal

Flow-Weighted Composite Sample

X X X X X X X X X

X X X X X X X

X

X

X

X

Flow-Weighted Composite Sample X X X X X X X X

X X X X X X X X X X

Source: From Friling, L., Pollut. Eng., 25, 36, 1993. With permission. X, analysis required. a

National pollutant discharge elimination system.

Both manual and automatic methods can be used to collect samples for the required analysis [8]. For manual sampling, the samples can be taken at fixed time intervals in individual bottles. After collection, a specific volume must be poured out of each bottle to form a flow-weighted composite. The exact volume must be calculated using the flow data taken when each bottle was filled. The advantage of manual sample collection is that, regardless of runoff amount, a fairly constant volume of sample is collected. This is because the flow-weighted composite is formed after the event and does not depend on calculations for runoff volume. Automatic storm water monitoring systems typically consist of a rain gauge, flowmeter, automatic sampler, and power source. The rain gauge measures on-site rainfall. The flowmeter measures the runoff water level and converts this level to a flow rate. In many systems, the flowmeter activates the sampler when user-specified conditions of rainfall and water level have been reached. Once activated, the sampler collects water samples by pumping the runoff water into bottles inside the sampler. Automatic storm water monitoring systems can form the flow-weighted composite sample automatically during the storm event, if there is sufficient storage capacity to accommodate variations in the runoff amount. Such automatic samplers are described in Refs. [8,16,17].

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1.4

Sampling Equipment

1.4.1

General Comments

This section discusses the advantages and disadvantages of sampling systems designed for the various sampling roles discussed above. Any component of a sampling device that is not normally present in the water body may affect the concentrations of determinants of interest in the water through three main effects: (a) by disturbance of physical, chemical, and biological processes and equilibria, (b) by contaminating the water with some parts of the sampler (e.g., organic compounds may leach out of plastic materials), and (c) by direct reactions between determinants and the materials of which the device is constructed (e.g., dissolved oxygen can react with copper, decreasing the oxygen concentration in the water). Another source of error may occur from processes within sample devices, such as the deposition of undissolved, solid materials on to the walls of a device, or sorption of chemicals to such surfaces. 1.4.2

Manual Sampling Systems

1.4.2.1 Simple Sampler for Shallow Water For many purposes, specially designed and installed sampling devices are not required. It often suffices simply to immerse a bottle in the water of interest, and this technique may be applicable also for some purposes in water-treatment plant. 1.4.2.2

Sampler for Large Quantities in Shallow Water

A system for the sampling and filtering of large quantities of surface seawater that is suitable for trace metal analysis is described in Ref. [35]. The water is brought aboard the ship via an all-Teflon pump and PFA tubing from a buoy deployed away from the vessel. The sample is delivered directly into a polycarbonate pressure reservoir and is subsequently filtered through a polycarbonate filter and in-line holder. Sampling systems based not on sample containers but on inlet tubes are commonly required in water-treatment and other plants, and are also employed in a number of applications for natural waters [5]. In some systems, the flow of sample through the inlet tube is achieved by the natural pressure differential, whereas in others the sample must be pumped, sucked (by vacuum), or pressurized (by a gas) through the tube. When dissolved gases and volatile organic compounds and possibly other determinants whose chemical forms and concentrations may be affected by dissolved gases are of interest, it is generally desirable to ensure that the sample is slightly pressurized to prevent gases coming out of solution. 1.4.2.3 Simple Sampler for Deepwater When it is necessary to sample from a particular depth in waters where the simple technique (see Section 1.2.2) cannot be used, special sample collection containers are available that can be lowered into the water on a cable to collect a sealed sample at the required depth. One of the simplest kinds of equipment with which to obtain samples from various depths is an empty weighted bottle closed with a stopper. This stopper is connected to the bottleneck by a rope, which can be used for releasing the stopper and opening the bottle at the desired depth (scoop bottle according to Meyer, Figure 1.4). However, for some sampling tasks, such as measuring dissolved gases, allowing the water flowing into the bottle to mix with the air inside the bottle is unacceptable.

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Cork

Sample bottle

Weight

FIGURE 1.4 Scoop bottle according to Meyer, a very simple sampler for various depths.

1.4.2.4 Deepwater Sampler (Not Adding Air to the Sample) A common tool for taking water samples from different depths is the standard water sampler according to Ruttner. The sampler, still open, is lowered by cable into the water. When it reaches the desired depth, a messenger is let down on the cable. Upon striking the standard water sampler, the messenger releases the closing mechanism and the lids of the sampling tube close. In some versions a separate cable is used to close the sampling bottle. The advantage is that no mixing of air with the sample will occur. But this system has the disadvantage that the inner surface of the sampling tube is in contact with water at all the depths through which the sampler travels on its way to the desired depth, and thus may introduce contaminants to the sample from the shallower layers through which it has passed. Figure 1.5 shows an apparatus from Hydrobios as an example of a sampler according to Ruttner. This version contains a thermometer ranging from 2 C to þ30 C, indicating the temperature of the sample; the temperature can easily be read through the plastic tube of the sampler. The water sample can be drawn off through the discharge cock in the lower lid for the various analyses. A similar version with a metal-free interior of the sampling tube for the determination of trace metals is also available. 1.4.2.5 Deepwater Sampler for Trace Elements (Allowing Air to Mix with the Sample) The Mercos Water Sampler from Hydrobios (Figure 1.6) is suitable for ultratrace metal analysis. It consists of a holder device and exchangeable 500 mL Teflon bottles as sampling vessels, which can be used down to 100 m water depth. All fittings are made of titanium. The sampler is attached to a plastic-coated steel hydrographic wire and lowered into the water in closed configuration in order to prevent sample contamination by surface water. Upon reaching the desired water depth, the sampler is opened by means of plasticcovered messengers. When the messenger hits the anvil, the silicone tubings spring up to allow water to flow in and air to leave the bottle. In the case of serial operation, a second messenger for release of the next sampler is set free at the same time. A disadvantage of the system is the contact between air and the water sample within the bottle.

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FIGURE 1.5 Standard water sampler according to Ruttner for various depths. Photograph from Hydrobios Apparatebau GmbH.

For the determination of trace elements in seawater the system sampling bottle ¼ storage bottle ¼ reaction vessel is used, so that the samples cannot be falsified by pouring from one vessel to another. In order to sterilize the bottles for microbiological investigations the Teflon bottles, along with coupling pieces and silicone tubings, can easily be taken from the holder.

FIGURE 1.6 MERCOS water sampler with two bottles which are lowered while closed and are opened at the desired depth. Photograph from Hydrobios Apparatebau GmbH.

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A modification of an inexpensive and easy-to-handle let-go system, a semiautomatic apparatus for primary production incubations at depths between 0 and 200 m, has been suggested [36]. The system is composed of a buoy, a nylon line, a fiberglass ballast weight, and about 15 sampling chambers. The entire volume of this apparatus is less than 80 dm3 and weighs about 7–8 kg. The sampling chambers sink with the ballast in an open position. When the line is stretched between the buoy at the surface and the ballast at the bottom, the chambers automatically enclose the water sample at the predetermined depth. The complete deployment of the apparatus takes less than 10 min. By an easy modification of the length of the line and/or the position of the chambers along it, sampling depth can be varied for repeated deployment over variable depth. The advantage of this system is that parallel samples from different depths can be obtained with relatively low costs and technical complexity. 1.4.3

Systems for Sampling the Benthic Boundary Layer at Different Depths

1.4.3.1 Deepwater (>50 m) Instrumented tripods with flowmeters, transmissiometers, optical backscatter sensors (OBS), in situ settling cylinders, and programmable camera systems have often been used in marine environments, for example, oceanographic studies of flow conditions and suspended particle movements in the bottom nepheloid layer [37,38]. These instruments were deployed to study suspended-sediment dynamics in the benthic boundary layer and were able to collect small water samples (1–2 L) at given distances from the seafloor. An instrumented tripod system (Bioprobe), which collects water samples and time-series data on physical and geological parameters within the benthic layer in the deep sea at a maximum depth of 4000 m, has been described [39]. For biogeochemical studies, four water samples of 15 L each can be collected between 5 and 60 cm above the seafloor. Bioprobe contains three thermistor flowmeters, three temperature sensors, a transmissiometer, a compass with current direction indicator, and a bottom camera system. 1.4.3.2 Shallow Water (