Tracking Genetically-Engineered Microorganisms

BIOTECHNOLOGY INTELLIGENCE UNIT 2 Janet K. Jansson • Jan Dirk van Elsas • Mark J. Bailey Tracking Genetically-Enginee...

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BIOTECHNOLOGY INTELLIGENCE UNIT

2

Janet K. Jansson • Jan Dirk van Elsas • Mark J. Bailey

Tracking Genetically-Engineered Microorganisms

BIOTECHNOLOGY INTELLIGENCE UNIT 2

Tracking Genetically-Engineered Microorganisms Janet K. Jansson Department of Biochemistry Stockholm University Stockholm, Sweden

Jan Dirk van Elsas Research Institute for Plant Protection Wageningen, The Netherlands

Mark J. Bailey National Environment Research Council Oxford, U.K.

LANDES BIOSCIENCE GEORGETOWN, TEXAS U.S.A.

EUREKAH.COM AUSTIN, TEXAS U.S.A.

TRACKING GENETICALLY-ENGINEERED MICROORGANISMS Biotechnology Intelligence Unit EUREKAH.COM LANDES BIOSCIENCE Designed by Judith Kemper Copyright ©2000 EUREKAH.COM All rights reserved. No part of this book may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Printed in the U.S.A. Please address all inquiries to the Publishers: EUREKAH.COM / Landes Bioscience, 810 South Church Street Georgetown, Texas, U.S.A. 78626 Phone: 512/ 863 7762; FAX: 512/ 863 0081 www.landesbioscience.com www.EUREKAH.COM

ISBN: 1-58706-009-4

Library of Congress Cataloging-in-Publication Data

Tracking genetically-engineered microorganisms / Janet K. Jansson, J.D. van Elsas, Mark J. Bailey. p.cm -- (Biotechnology intelligence unit) Includes bibliographical references and index. ISBN 1-58706-009-4 (alk. paper) I. Recombinant microorganisms -- Ecology. 2. Recombinant microorganisms -- Identification. I. Jansson, Janet K. II. Elsas, J.D. van (Jan D.), 1951- III. Bailey, Mark J. IV. Series. QR 100.T73 2000 660.6'5--dc21

99-056991

CONTENTS 1. Problems in Detecting Dormant (VBNC) Cells, and the Role of DNA Elements in This Response ......................................................... 1 James D. Oliver 1.1. The Viable but Nonculturable State ............................................. 1 1.2. Assays Used to Determine Viability .............................................. 2 1.3. Characteristics of Cells in the VBNC State ................................... 3 1.4. Inducing Factors ............................................................................ 4 1.5. Bacteria Known to Enter the VBNC State .................................... 5 1.6. Resuscitation from the VBNC State .............................................. 5 1.7. In situ Evidence of the VBNC Response ....................................... 7 1.8. Use of PCR to Detect Cells in the VBNC State ............................ 9 1.9. Effect of Plasmids on Entry into the VBNC State ........................ 9 1.10. Significance of the VBNC State in the Release of Genetically Modified Bacteria ................................................. 12 1.11. Conclusion ................................................................................... 13 2. The Use of Antibiotic Resistance Gene Markers for Studying Bacterial Populations in Natural Environments ............. 17 Sharon Egan and Elizabeth M.H. Wellington 2.1. Introduction ................................................................................. 17 2.2. Cultivation Based Assays ............................................................. 18 2.3. Direct Molecular Monitoring Methods ...................................... 20 2.4. Choice of Antibiotic Resistance Gene Markers .......................... 20 2.5. Cross-Resistance .......................................................................... 21 2.6. Marking of Bacteria with Antibiotic Resistance Genes .............. 21 2.7. Use of Antibiotic Resistance Genes to Monitor Gene Transfer in Soil ................................................ 22 2.8. Ethical Concerns .......................................................................... 23 2.9. Conclusion ................................................................................... 24 3. Extraction and Analysis of Microbial Community Nucleic Acids from Environmental Matrices ................................................................ 29 Jan Dirk van Elsas, Kornelia Smalla and Christoph C. Tebbe 3.1. Introduction ................................................................................. 29 3.2. Extraction of Microbial Cells from Environmental Matrices .... 33 3.3. Cell Disruption ............................................................................. 35 3.4. Extraction and Purification of DNA ........................................... 36 3.5. Analysis of DNA and Detection of Specific Sequences .............. 38 3.6. Extraction and Purification of RNA ........................................... 39 3.7. Detection of RNA ......................................................................... 40 3.8. Quantification of Specific Targets in Environmental Nucleic Acids ................................................. 42 3.9. Conclusion ................................................................................... 44

4. Detection of Bacteria by Their Intrinsic Markers ................................. 53 Éva Tas and Kristina Lindström 4.1. Introduction ................................................................................. 53 4.2. Brief Description of PCR-Based Fingerprinting Methods ......... 54 4.3. Development of Specific Hybridization Probes and PCR Primers .......................................................................... 54 4.4. Case Study: Specific Detection of Rhizobium galegae ................ 56 4.5. Future Prospects .......................................................................... 61 4.6. Conclusion ................................................................................... 63 5. Luminescence-Based Microbial Marker Systems and Their Application in Microbial Ecology ......................................... 69 James I. Prosser, Antonio J. Palomares, Matti T. Karp, Philip J. Hill 5.1. Introduction ................................................................................. 69 5.2. Lux-Based Systems ....................................................................... 69 5.3. Luc-Based Systems ....................................................................... 72 5.4. Methodology ................................................................................ 74 5.5. Applications ................................................................................. 76 5.7. Conclusion ................................................................................... 82 6. The Use of the GUS Reporter System to Study Molecular Aspects of Interactions Between Bacteria and Plants ......................................... 87 M. Lambrecht, A. Vande Broek and Jos Vanderleyden 6.1. Introduction ................................................................................. 87 6.2. The Escherichia coli gusA gene ..................................................... 88 6.3. The GUS Reporter system ........................................................... 89 6.4. Application of the GUS Reporter System in Studies of Plant-Bacterium Interactions ................................ 93 6.5. Conclusion ................................................................................... 97 7. Using Green Fluorescent Protein (GFP) as a Biomarker or Bioreporter for Bacteria ......................................... 101 J. R. Stoltzfus, Janet K. Jansson and Frans J. de Bruijn 7.1. Introduction ............................................................................... 101 7.2. Properties of Wild-type GFP ..................................................... 101 7.3. Improving GFP Fluorescence .................................................... 102 7.4. Detection of GFP ....................................................................... 104 7.5. GFP as a Biomarker in Bacteria ................................................. 105 7.6. GFP as a Bioreporter in Bacteria ............................................... 111 7.7. Conclusion ................................................................................. 112

8. Monitoring Persistence and Risk Assessment Following the Field Release of Pseudomonas fluorescens SBW25EeZY6KX ........ 117 Mark J. Bailey, Tracey M. Timms-Wilson, Richard J. Ellis, Ian P. Thompson and Andrew K. Lilley 8.1. Introduction ............................................................................... 117 8.2. Construction of P. fluorescens SBW25EeZY6KX and Detection and Monitoring Methods ................................. 118 8.3. Case Study of the Persistence of P. fluorescens SBW25EeZY6KX in Sugar Beet Crops ........... 118 8.4. Survival of Recombinant in Bulk Soil under Glasshouse Conditions ................................................... 119 8.5. Field Sites, Treatment and Sampling of Plants and Soil .......... 120 8.6. Survival of GMM on Over-Wintering Sugar Beet Secondary Growth, Resown Crop Plants and Volunteer Weeds ............... 120 8.7. Would Persistence of a Modified Indigenous Strain in the Soil Environment Pose a Threat? .................................... 122 8.8. Conclusion ................................................................................. 123 9. Use of luc-Tagged Genetically Modified Microorganisms (GMMs) to Study Rhizobial Ecology in Soil Columns, Field Lysimeters and Field Plots ........................................................... 127 Christoph C. Tebbe 9.1. Introduction ............................................................................... 127 9.2. Construction and Properties of GMM Strains L1 and L33 ..... 128 9.3. Experimental Design: What Can Be Achieved with Each System? ...................................................................... 129 9.4. Techniques and Consequences of GMM Soil Inoculations ..... 132 9.5. Survival and Spread of GMMs in Soil Columns, Field Lysimeters and Field Plots ................................................ 133 9.6. Conclusion ................................................................................. 135 10. The Field Release and Monitoring of Rhizobial Strains Marked with lacZ and Mercury Resistance Genes ............................................ 139 Viviana Corich, Alessio Giacomini, Elena Vendramin, Patrizia Vian, Milena Carlot, Andrea Squartini and Marco P. Nuti 10.1. Introduction ............................................................................... 139 10.2. Designing the Markers ............................................................... 139 10.3. Construction of GMM Strains 1110, 1111 and 1112 ............... 140 10.4. Microcosm Studies of GMMs ................................................... 140 10.5. Field Trial I; Fate and Impact of an Allochtonous GMM ........ 141 10.6. Field Trial II; Fate and Impact of a GMM Native to the Site .................................................................................... 143 10.7. Conclusion ................................................................................. 144

11. The Field Release and Monitoring of GUS-Marked Rhizobial Strain CT0370 ........................................... 145 Penny R. Hirsch, Tom A. Mendum, Alfred Pühler and Werner Selbitschka 11.1. Introduction .............................................................................. 145 11.2. Construction of GMM Strain CT0370 .................................... 145 11.3. Detection of Strains CT0370 and RSM2004 ........................... 146 11.4. Inoculant Preparation .............................................................. 147 11.5. Field Release .............................................................................. 147 11.6. Screening for pSym Acquisition by CT0370 in the Field ....... 148 11.7. Plasmid Transfer from RSM2004 to CT0370 .......................... 148 11.8. Conclusion ................................................................................ 149 12. Regulatory Aspects ................................................................................ 153 Kersti Gustafsson 12.1. Introduction .............................................................................. 153 12.2. Historical Aspects of the Regulation of Deliberate Releases of GMOs ............................................... 153 12.3. The European Union and the European Free Trade Association .............................. 155 12.4. Canada ....................................................................................... 156 12.5. Switzerland ................................................................................ 157 12.6. United States ............................................................................. 157 12.7. Risk Assessment ........................................................................ 158 12.8. Concern about the Insertion of Genes Coding for Antibiotic Resistance .......................................................... 159 12.9. Concern about the Insertion of Genes Coding for Mercury Resistance ............................................................. 159 12.10. Concern on the Use of Hazardous Chemicals in Connection with the Deliberate Release of a GMM ........... 159 12.11. Ethical Aspects .......................................................................... 160 12.12. Information on Field Releases and Products .......................... 160 Index ....................................................................................................... 163

EDITORS Janet K. Jansson Department of Biochemistry Stockholm University Stockholm, Sweden Chapter 7 Jan Dirk van Elsas Research Institute for Plant Protection Wageningen, The Netherlands Chapter 3 Mark J. Bailey National Environment Research Council Institute of Virology and Environmental Microbiology Oxford, U.K. Chapter 8

CONTRIBUTORS A. Vande Broek F.A. Janssens Laboratory of Genetics Kardinaal Mercierlaan 92 Heverlee, Belgium Chapter 6

Sharon Egan Department of Biological Sciences University of Warwick Coventry, U.K. Chapter 2

F.J. de Bruijn MSU-DOE Plant Research Laboratory Department of Microbiology NSF Center for Microbial Ecology Michigan State University East Lansing, Michigan, U.S.A. Chapter 7

R.J. Ellis Molecular Microbial Ecology Laboratory NERC, Institute of Virology and Environmental Microbiology Oxford, U.K. Chapter 8

Milena Carlot Dipartimento di Biotecnologie Agrarie Università di Padova Legnaro, Italy Chapter 10

Alessio Giacomini Dipartimento di Biotecnologie Agrarie Università di Padova Legnaro, Italy Chapter 10

Viviana Corich Dipartimento di Biotecnologie Agrarie Università di Padova Legnaro, Italy Chapter 10

Kersti Gustafsson Microbiologist/Ecotoxicologist National Chemicals Inspectorate Solna, Sweden Chapter 12

Philip J. Hill School of Biological Sciences University of Nottingham Sutton Bonington Campus Sutton Bonington, Leicestershire, U.K. Chapter 5

Marco P. Nuti Dipartimento di Chimicae Biotecnologie Agrarie Università di Pisa Pisa, Italy Chapter 10

Penny R. Hirsch Soil Science Department IACR-Rothamsted Harpenden, U.K. Chapter 11

James D. Oliver Professor of Biology Director, Interdisciplinary Biotechnology Program University of North Carolina Charlotte, North Carolina, U.S.A. Chapter 1

Matti T. Karp Department of Biotechnology University of Turku Tykistokatu Turku, Finland Chapter 5 M. Lambrecht F.A. Janssens Laboratory of Genetics Kardinaal Mercierlaan 92 Heverlee, Belgium Chapter 6 A.K. Lilley Molecular Microbial Ecology Laboratory NERC, Institute of Virology and Environmental Microbiology Oxford, U.K. Chapter 8 Kristina Lindström Department of Applied Chemistry and Microbiology Division of Microbiology Viikki Biocenter University of Helsinki Helsinki, Finland Chapter 4 Tom A. Mendum Soil Science Department IACR-Rothamsted Harpenden, U.K. Chapter 11

Antonio J. Palomares Departamento de Microbiologia y Parasitologia Universidad de Sevilla Sevilla, Spain Chapter 5 James I. Prosser Department of Molecular and Cell Biology University of Aberdeen Institute of Medical Sciences, Foresterhill Aberdeen, Scotland Chapter 5 Alfred Pühler Department of Genetics University of Bielefeld Bielefeld, Germany Chapter 11 Werner Selbitschka Department of Genetics University of Bielefeld Bielefeld, Germany Chapter 11

Kornelia Smalla Institut fuer Biochemie und Pflanzenvirologie, Biologische Bundesanstalt Braunschweig, Germany Chapter 3 Andrea Squartini Dipartimento di Biotecnologie Agrarie Università di Padova Legnaro, Italy Chapter 10 J.R. Stoltzfus MSU-DOE Plant Research Laboratory Department of Microbiology NSF Center for Microbial Ecology Michigan State University East Lansing, Michigan, U.S.A. Chapter 7 Éva Tas Department of Biosciences Division of Genetics Viikki Biocenter University of Helsinki Helsinki, Finland Chapter 4 Christoph C. Tebbe Institut fuer Agraröecologie, FAL Braunschweig, Germany Chapters 3, 9 Ian P. Thompson Molecular Microbial Ecology Laboratory NERC, Institute of Virology and Environmental Microbiology Oxford, U.K. Chapter 8

Jos Vanderleyden F.A. Janssens Laboratory of Genetics Kardinaal Mercierlaan 92 Heverlee, Belgium Chapter 6 Elena Vendramin Dipartimento di Biotecnologie Agrarie Università di Padova Legnaro, Italy Chapter 10 Patrizia Vian Dipartimento di Biotecnologie Agrarie Università di Padova Legnaro, Italy Chapter 10 Elizabeth M.H. Wellington Department of Biological Sciences University of Warwick Coventry, U.K. Chapter 2 Tracey M. Timms-Wilson Molecular Microbial Ecology Laboratory NERC, Institute of Virology and Environmental Microbiology Oxford, U.K. Chapter 8

PREFACE

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nvironmental microbiology is currently one of the most rapidly expanding areas of scientific research. Impetus for advanced investigations has been provided by the development and application of molecular techniques that facilitate the identification, characterization and monitoring of microbes. These advances now allow detailed investigations, developed in the laboratory, to be undertaken in the natural environment. Such studies confirm the remarkable biological diversity represented by microorganisms from their basic genetic structure to the regulated communications that occur within and between communities contributing to ecosystem function. However, while it is apparent that microorganisms constitute the greater part of the planet’s biomass, and are central in maintaining the biosphere, we remain essentially ignorant of a great majority of the functions or processes they undertake. One of the limiting factors in the study of their ecology, even within communities or populations, is that of scale. For instance, for soil it is very hard to assess microbes and their activities at the level of each individual pore where microbial soil inhabitants occur. Highly sensitive and specific tools are, thus, required for such detailed investigations. However, there is a paradox. Until a greater knowledge of the genetic and metabolic diversity within the microbial environment is obtained it remains difficult to investigate these complex communities or design relevant experiments that target the role of individuals or specific genes and their products. This situation is currently changing at an ever increasing rate. In this volume we have attempted to bring together a series of reviews of the approaches taken to study the ecology and functional activity of individual microbial cells and populations in environmental habitats, with a special focus on the use of marker/reporter genes for monitoring release strains. Traditionally, microbial ecology has been limited to studies of microbial processes, such as cycling of nitrogen or carbon, occurring by uncharacterized species in a “black box” scenario. For example, denitrification, nitrification and nitrogen fixation processes have been assessed by analyzing the specific nitrogen forms appearing as a result of these processes. On the other hand, the study of particular taxa was limited to those for which suitable cultivation methods and laboratory growth media had been devised. However, we now know that the majority of microorganisms in nature are not capable of growing on the available media under standard laboratory conditions. Moreover, in some instances bacteria, that have been successfully cultivated in the laboratory, may lose the ability to grow on laboratory media after introduction into the environment, presumably as a result of a stress response. These bacteria may still be viable or metabolically active in the environment and this apparently recalcitrant state must be considered when these organisms are studied. The

ability to persist and form a “resting stage” may be ecologically significant in niche exploitation. The phenomenon of viable but non-culturable (VBNC), as described for Vibrio spp, is discussed in Chapter 1. In the 1980s the need to identify and track specific microorganisms in environmental samples was highlighted by the requirement for risk assessment of genetically modified microorganisms (GMMs) that were to be deliberately released into natural settings. The current status of regulatory requirements for GMMs is discussed in Chapter 12. For risk assessment of GMMs it is no longer possible to rely purely on traditional monitoring methods since the specific GMM has to be distinguished from its parent, or wild type, strain that could also inhabit the same ecosystem. Based on the requirement for specific monitoring methods, a variety of molecular tools have been developed to unambiguously track GMMs in the environment. These tools and examples of their applications for tracking GMMs in field studies are described in this book. The molecular tools range from nucleic acid-based monitoring methods (Chapter 3) to the use of marker genes (biomarkers) with distinct phenotypes that serve as tags for identification of specific microbes in the laboratory (Chapters 5-7) and in the natural environment (Chapters 8-11). Following the established method of isolating defined strains using selective bacteriological media, nucleic acid-based molecular tools were the first to be applied for the purpose of directly tracking GMMs in environmental samples. This methodology was dependent on the ability to isolate nucleic acids from the environment, and a series of methods were developed for the extraction of DNA and more recently RNA, from complex matrices such as soil. These methods have been continually refined since the first published examples (reviewed in Chapter 3). As the technology improves, further optimizations in sensitivity and specificity can be expected. A variety of nucleic acid extraction methods were originally optimized for the different environmental sample types under analysis. These extraction and analytical methods have now been simplified and applied as essentially standard protocols for tracking GMMs in any environment. In particular, the application of polymerase chain reaction amplification (PCR) has revolutionized nucleic acid-based tracking methods by substantially increasing the sensitivity of detection over nucleic acid hybridization methods (Chapter 3). A biomarker can be defined as a DNA sequence, introduced into an organism, which confers a distinct genotype or phenotype to enable monitoring in a given environment. In some cases, an intrinsic marker is sufficient for monitoring a particular bacterial species. An intrinsic marker is a nonintroduced DNA sequence or a natural genotype that serves as a signature for a particular organism or group of organisms. Intrinsic markers are further described in Chapters 2 and 4. Chapter 2 discusses antibiotic resistance as a special type of intrinsic marker. Usually, intrinsic markers are not sufficiently specific for tracking of GMMs, since they are also present in the parent, or “wild-type”, strain, or they may be prevalent in the environment under study. For some applications, a reporter gene (bioreporter) can be used. A bioreporter is a gene encoding an easily detectable phenotype that can be used to measure gene expression when

fused to appropriate promoter target sequences. The choice of biomarker or bioreporter depends on the particular strain, the environment studied and the questions to be addressed. Several of the most promising monitoring methods and examples of their application are described in detail in Chapters 5-7 of this book. In Chapters 8-11, examples are given of the use of these biomarkers to track specific GMMs after release during field trial experiments. Chapter 12 outlines regulatory considerations for risk assessment of GMMs before field release. Figure 1 gives a schematic overview of the use of biomarkers/bioreporters in GMM releases. The use of any type of marker/reporter is seen by many as being essential in assessing the environmental fate of release organisms. Each chapter of this book is authored by at least one participant of the MAREP Concerted Action of Scientists. MAREP is an acronym for “Marker/ Reporter genes in Microbial Ecology” and is comprised of 26 scientists from 11 different countries. The MAREP Concerted Action is funded by the European Commission, Directorate-General XII, and addresses issues related to the use of marker and reporter genes in microbial ecology research. This is of particular importance in relation to the precise monitoring of GMM presence, persistence and metabolic activity in the environment. More information about the MAREP Concerted Action can be found in the following Web site: http://www.biokemi.su.se/marep/marep.html.

We intend this book to be of relevance for all those concerned with studies of the environmental fate of genetically modified or unmodified microorganisms. In particular, this volume will be of value to researchers developing organisms intended for release, and to representatives of regulatory agencies concerned with guiding experimental or commercial applications. This volume provides an up-to-date collection of data on the development, use and assessment of biomarkers and bioreporters for the study of bacterial function in the natural environment. We would like to extend our thanks to all the authors for providing the necessary text for this publication, and for the patience and understanding they have shown during the editing process. We would also like to thank the editorial staff for their support. Janet K. Jansson Jan Dirk van Elsas Mark J. Bailey

CHAPTER 1

Problems in Detecting Dormant (VBNC) Cells, and the Role of DNA Elements in This Response James D. Oliver

1.1. The Viable but Nonculturable State

M

icrobial ecologists have long recognized that large proportions of microbial populations inhabiting natural habitats appear to be nonculturable. Indeed, plate counts of bacteria in soil, rivers and oceans typically indicate that far less than 1% of the total bacteria observed by direct microscopic examination can be grown on culture media. It has also long been known that certain portions of bacterial populations in natural environments seem to “disappear” during certain seasons, only to “reappear” at other times. We now understand that at least part of the explanation for these observations is not due to seasonal die-off of the cells, but to their entry into what is most commonly called the “viable but nonculturable” state.1 A bacterial cell in the viable but nonculturable (VBNC) state may be defined as one which fails to grow on the routine bacteriological media on which it would normally grow and develop into a colony, but which is in fact alive and metabolically active. Bacteria enter into this “dormant” state in response to one or more environmental stresses which might otherwise be ultimately lethal to the cell. Thus, the VBNC state should be considered a means of cell survival. Eventually, when the inducing stress is removed, these cells are able to emerge from the VBNC state, and again become culturable on routine media. The typical VBNC response is illustrated in Figure 1.1, which shows the response of the human pathogen, V. vulnificus, to exposure to low temperature (5˚C). Such a temperature is below that at which this aquatic bacterium can grow and, if it were not for the VBNC response, is a temperature which would eventually lead to death of the population. As is evident from Figure 1.1, cells lose their ability to be cultured (shown by the open squares) in a rather linear manner, eventually reaching a point where platings suggest a total lack of any living cells. However, whereas death of a bacterial population generally leads to lysis of the cells and loss of cell structure, direct examination of cells entering the VBNC state indicates that the cells remain intact (as shown by the open circles of Fig. 1.1). Such cells could, of course, have died, but simply not undergone lysis. The primary evidence that such cells are alive, even if nonculturable, is from data obtained when one of the “direct viability” assays is applied to such cultures. These assays (described below) allow the direct

Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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Fig. 1.1. Entry of V. vulnificus into the VBNC state in an artificial sea water microcosm at 5oC. Shown are plate counts (❏) on HI agar in cfu/ml, total cell counts (◆) by the acridine orange staining method, and direct viable counts (❍) by the substrate responsive method using yeast extract and nalidixic acid. Reprinted with permission from: Whitesides MD, Oliver JD. Appl Environ Microbiol 1997; 63:1002-1005.

determination of the viability of individual cells in a population, without the need for culture. As seen in Figure 1.1 (open circles), such assays often indicate that a large proportion of the apparently dead population is indeed alive.

1.2. Assays Used to Determine Viability Several techniques now exist for demonstrating that a cell is viable without requiring growth in or on a bacterial medium. These fall roughly into two categories: those which demonstrate an active electron transport chain, and those which demonstrate the ability of cells in the VBNC state to undergo limited growth, if not cellular division. In the former, the most common indicators are reduction of INT (p-iodonitrotetrazolium violet) or of CTC (5-cyano-2,3-ditolyl tetrazolium chloride). INT is an electron acceptor which diverts electrons from an active electron transport chain, thus becoming reduced to INT-formazan. Whereas INT is soluble and colorless, INT-formazan is insoluble and has a purple to black color. Thus, a cell which is undergoing metabolism using an electron transport chain (and therefore presumably is viable) will reduce INT to INT-formazan. Because the electron transport chain is in the bacterial inner membrane, this reduction leads to the development of dark spots in the membrane, representing sites of precipitation of the INT-formazan. These spots are readily detected using the light microscope. Through a similar action, reduction converts CTC to CTC-formazan, which fluoresces when irradiated by ultraviolet light. The result is red-fluorescent cells which are readily detected and differentiated

Detection of VBNC Cells

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from non-fluorescing (and presumably non-viable) cells by epifluorescent microscopy. Both methods are described in Oliver.1 The most common method, originally developed by Kogure et al,2 for demonstrating viability other than through an active electron transport chain is that involving the addition to VBNC cells of small amounts of nutrient, usually in the form of yeast extract, along with nalidixic acid, an inhibitor of cell septation. Cells which are viable and nutrient responsive metabolize the yeast extract, and begin to elongate. The antibiotic prevents their cell division, however, resulting in elongated cells (Fig. 1.2). Such cells, which clearly are viable, are easily distinguished from nonviable, and therefore non-elongated, cells.1 Luminescence-based detection methods3,4 and flow cytometry5 may prove of great value in detecting the activity of cells as they enter into, and resuscitate from, the VBNC state. Other technologies, such as the use of fluorescently-tagged polyclonal or monoclonal antibodies6 are also essential for the detection of VBNC cells in the environment.

1.3. Characteristics of Cells in the VBNC State Cells entering the VBNC state generally undergo a reduction in size. In the case of V. vulnificus, for example, whereas log phase (actively growing) cells might be 3 mm long, those in the VBNC state are typically 0.6 µm in diameter. During this size reduction, significant changes in membrane structure, protein composition, ribosomal content, and possibly even DNA arrangement are experienced. Again using V. vulnificus as an example,1 we have found rapid and dramatic decreases in the levels of synthesis of DNA, RNA, and protein when these cells are exposed to a temperature downshift to 5˚C (Fig. 1.3). However, such decreases do not mean that all synthesis has ceased. Indeed, protein synthesis appears to be essential for entry into this state, and under these conditions V. vulnificus produces some 40 new proteins not seen during growth at “normal” temperatures.7 At the same time, dramatic changes in membrane fatty acid composition,8 and decreases in nutrient transport and respiration rates have generally been reported to occur as cells enter this dormant state.

Fig. 1.2. Elongation of viable cells following addition of yeast extract and nalidixic acid by the method of Kogure et al.2 Nonviable cells remain as small, coccoid cells.

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Fig. 1.3. Macromolecular synthesis in V. vulnificus during entry into the VBNC state at 5˚C. Cells were assayed for protein (■ ), DNA (❏ ), and RNA (◆) synthesis. Reprinted with permission from: Oliver JD. In: Kjelleberg S, ed. Starvation in Bacteria. New York: Plenum Press 1993:239-272.

Cell wall synthesis, or at least metabolism of the constituents of these structures, also apparently continues, as addition of penicillin (an inhibitor of cell wall synthesis) to VBNC cells has generally been observed to result in rapid cell death.1 Most studies have observed that, if a cell entering the VBNC state harbors plasmids (extrachromosomal DNA elements which are able to control a variety of generally nonessential cell functions), then these plasmids are retained. This finding may prove to be highly relevant to the VBNC state of genetically modified cells released to the environment, as will be discussed later in this chapter. In contrast, it is becoming increasingly apparent that (possibly even major) changes in the cells’ chromosomal DNA may be occurring as cells enter the VBNC state.1 This aspect may also be critical to release studies, as these changes bring into question the ability to employ such powerful molecular techniques as the polymerase chain reaction (PCR) to detect these otherwise undetectable cells. This concern is also dealt with later in this chapter.

1.4. Inducing Factors Different bacteria are known to enter the VBNC state in response to different factors, all of which a cell would normally encounter in natural environments. These include such stresses as elevated or reduced temperature, elevated or reduced osmotic (e.g., salt) concentrations, nutrient starvation, levels of oxygen, and even certain intensities of light.1 In all cases, the inducers of the VBNC state appear to be environmental factors which are potentially injurious to a given bacterial species. For example, while entry into the VBNC state by V. vulnificus (optimum temperature of 37˚C) is induced by low (< 10˚C) temperature incubation,9 the reverse is true for Pseudomonas fluorescens. One strain of this bacterium, which prefers low temperatures, enters the VBNC state at 37˚C.10


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The time required for cells to enter the VBNC state varies markedly with the bacterium and the inducing conditions. Reports of months being required are not uncommon, while others have reported days for the same bacteria. In general, times of a few days to a month seem typical. One factor which has been shown to have a dramatic effect on the time required for lab-grown cells to become nonculturable is the “age” of the cells. We have shown that, whereas V. vulnificus cells from the logarithmic phase of growth generally require less than 10 days to become completely nonculturable at 5˚C, those taken from the stationary phase require over a month. Indeed, a direct correlation between time required to become nonculturable and the age of the population has been demonstrated.11

1.5. Bacteria Known to Enter the VBNC State At least 30 species contained in16 genera of mostly gram-negative bacteria have been demonstrated to enter the VBNC state12 (Table 1.1). That such a variety of bacterial genera demonstrate this response suggests that it represents a wide-spread physiological response to adverse environments designed to allow the cell to survive potentially lethal environmental stresses. Whether this stress response provides cross protection against other stresses has not been reported.

1.6. Resuscitation from the VBNC State For the VBNC response to represent a true survival response it must be possible for the cells to exit this dormant state and return to a fully active and culturable state. Such a reversal in physiology is termed resuscitation, and often is triggered simply by the removal of the stress which initially induced the VBNC response. In V. vulnificus, for example, exit from the low temperature-induced VBNC state is triggered by a temperature upshift (e.g., from 5˚C to room temperature). After such a shift, culturable cells rapidly (typically within 8 hours) begin to appear, and population levels approximately equaling the original levels are generally observed within 12-24 hours (Fig. 1.4). During this time, the small coccoid cells which result as the cells enter the VBNC state are replaced by rods typical in size for V. vulnificus.13

Table 1.1. Bacteria described to enter the VBNC state Aeromonas salmonicida Agrobacterium tumefaciens Campylobacter jejuni Enterobacter aerogenes Enterococcus faecalis Escherichia coli (including EHEC strains) Helicobacter pylori Klebsiella pneumoniae Lactobacillus plantarum Legionella pneumophila Micrococcus luteus M. varians Pasteurella piscida Pseudomonas aeruginosa P. fluorescens P. putida P. syringae

Salmonella enteritidis S. typhimurium Shigella dysenteriae S. flexneri S. sonnei Vibrio anguillarum V. campbellii V. cholerae V. fischeri V. harveyi V. mimicus V. natriegens V. parahaemolyticus V. proteolytica V. vulnificus (biotypes 1 and 2)

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Fig. 1.4. Changes in culturable cell (plate) counts and cell morphology during temperature downshift to 5˚C and subsequent resuscitation of the nonculturable cells by incubation at ca. 22˚C. Reprinted with permission from: Nilsson L et al. J Bacteriol 1991; 173:5054-5059.

While entry into a VBNC state has been described by many researchers and for many different bacterial species, demonstrating resuscitation for these cells has not always been a simple matter. Indeed, while some bacteria like V. vulnificus can be resuscitated by a simple reversal of the inducing stress, in others it has been quite difficult to show. We now realize that resuscitation may be an extremely complex event, one which may be quite difficult to demonstrate in the lab. In the case of L. pneumophila, simple addition of nutrients to the cells (which enter the VBNC state in response to nutrient deprivation) does not reverse the dormancy. It was found, however, that the addition of certain amoebae, which are natural hosts to this bacterium in the aquatic environment, does allow resuscitation of this causative agent of Legionnaire’s disease.14 Other problems also exist in demonstrating resuscitation from the VBNC state. It has been difficult to overcome the argument that what was being observed in the name of “resuscitation” was, in fact, regrowth of a few culturable cells which had escaped detection during plating of the population under study. However, we have recently presented very strong evidence that, at least in the case of V. vulnificus, true resuscitation does occur.15 Our studies employed extensively diluted populations of VBNC cells in which it was statistically impossible that any culturable cells were present. Resuscitation of these populations occurred at such a rapid rate that, if it were due to regrowth of culturable cells, they would have to have had a doubling time of approximately 6 minutes. This is clearly an impossible generation time for cells incubated at a suboptimal temperature without nutrients or aeration (Fig. 1.5). We also observed that the presence of nutrients appears to inhibit (but not kill) VBNC cells, and this may be the reason such cells are “nonculturable” when plated onto the high organic nutrient media routinely employed for bacterial culture.


7

Fig. 1.5. Time required for resuscitation of VBNC V. vulnificus cells. Cells from a VBNC microcosm (< 3.3 x 101 cfu/ml) were shifted to room temperature and aliquots removed at hourly intervals and plated onto HI agar. Reprinted with permission from: Whitesides MD, Oliver JD. Appl Environ Microbiol 1997; 63:1002-1005.

1.7. In Situ Evidence of the VBNC Response This review has so far described laboratory findings regarding the VBNC state. One can validly question whether this response actually occurs in natural environments. We have conducted studies employing membrane diffusion chambers which clearly demonstrated the entry of V. vulnificus into the VBNC state when the estuarine waters into which they were submerged were cold.16 Similarly, when cells in the VBNC state were placed into chambers which were suspended into warm (summer) estuarine waters, resuscitation from the VBNC state occurred rapidly and to levels approximating those of the original population (Fig. 1.6). Thus, it is likely that our inability to isolate V. vulnificus during the winter months may be accounted for by entrance into the VBNC state, and that recovery from this state in natural environments may result from a temperature upshift. Taking these observations to their logical conclusions, it seems quite likely that the VBNC response may be a significant source of the variations seen in natural populations as they undergo various environmentally-induced stresses, and for inability to culture many of the microorganisms observed in natural environments. Indeed, Wilson and Lindow17 examined the culturability of P. syringae inoculated onto leaf surfaces, and found that up to 75% of the cells became nonculturable after 80 hours.

8


Fig. 1.6. Entry into (Fig. 1.6A) and resuscitation from (Fig. 1.6B) the VBNC state by V. vulnificus placed into membrane diffusion chambers in estuarine waters of the coast of North Carolina. A. Cells were placed into water at a temperature of 10-15˚C. Shown are plate counts (❍), total direct counts (●), and direct viability assays (❏). B. Plate counts of both the encapsulated (●) and non-encapsulated (❍) forms of V. vulnificus induced into the VBNC state in the laboratory at 5˚C. Four days after entry into the VBNC state (day 11), resuscitation of the cells is seen when placed into estuarine waters at a temperature of 16-19˚C. Reprinted with permission from: Oliver JD, Hite MF, McDougald D et al. Appl Environ Microbiol 1995; 61:2624-2630.


9

1.8. Use of PCR to Detect Cells in the VBNC State Alternate methods for detecting cells present in the VBNC state must be explored or developed. One powerful molecular method which would seem to be ideal for this purpose is the polymerase chain reaction (PCR). This method allows the specific DNA of the target bacterial population (in this case, the released GMOs) to be amplified, to the exclusion of that from other cells, to an extent that the amplified DNA can easily be detected, and thus the presence of the target cells determined. The use of PCR technology to amplify DNA from VBNC cells would seem to be an ideal match. However, in our earliest studies on the use of PCR methodology to detect cells of V. vulnificus present in the VBNC state, we found that as many as 500 times more VBNC cells had to be present to obtain an amplification signal as compared to culturable cells. Even when the DNA was extracted prior to amplification, 300 times or more DNA had to be present.18 In subsequent studies, we employed a variation on the PCR method, termed “randomly amplified polymorphic DNA” (RAPD) PCR analysis. To our surprise, when cells of V. vulnificus were induced into the VBNC state by incubation at 5˚C, our ability to detect amplification gradually decreased, until all bands were lost within a few days (Fig. 1.7). When the cells were subsequently allowed to resuscitate, RAPD-PCR amplification again occurred.19 The same observation has also been made by Bej et al20 with V. vulnificus. We surmise that, on incubation at the lowered temperature, the cells begin to synthesize cold shock-induced proteins, which prevent amplification of the DNA either by modifying the supercoiling of, or physically blocking the DNA polymerase from binding to, the target DNA. A similar result was observed when we used RAPD-PCR in an attempt to amplify cells of enterohemorrhagic E. coli as they entered the VBNC state.21 Even more dramatic was our finding that the RAPD-PCR signals we normally observed in V. vulnificus were lost within hours when the cells were deprived of nutrients.19 Resumption of amplification was observed only when nutrient was added back to the system, similar to the response seen when cells were resuscitated from the VBNC state. In all cases, loss of PCR amplification could be prevented when the cells were incubated in the presence of chloramphenicol, a protein synthesis inhibitor. This finding again suggests that protein synthesis is essential for cells to enter these stress-induced response states. The results of these studies bode poorly for the use of PCR technology as an alternative method for the detection of GMOs released to the environment which, for whatever reason, enter the VBNC state.

1.9. Effect of Plasmids on Entry into the VBNC State To make the situation potentially worse, it now appears that our knowledge of what induces the VBNC state in various bacteria must be re-examined, taking into account whether or not the cells under investigation harbor extrachromosomal plasmids. We have reported10 that Pseudomonas fluorescens cells enter the VBNC state when incubated in sterile river water at 37˚C, but not at 5˚ or 25˚C (Fig. 1.8A). In rather remarkable contrast, when this same strain harbored a plasmid (pFAC510), the cells entered the VBNC state at 5˚C, but not at 37˚C (Fig. 1.8B). The same result was seen for P. syringae, with and without an entirely different plasmid (pQF70/19H). Most recently, we have observed similarly dramatic plasmid effects when P. fluorescens is incubated in soil microcosms (Fig. 1.9). The fact that plasmids may have such a dramatic affect on culturability has obvious and significant implications for the monitoring of GMOs intended for environmental release. Indeed, the plasmid-induced “temperature reversal” observed in our studies suggests that such cells must be thoroughly characterized as to their plasmid content, with the realization that plasmid size and copy number may affect the temperature at which they enter the VBNC state.

10


Fig. 1.7. Randomly amplified polymorphic (RAPD) DNA analysis of V. vulnificus cells induced into the VBNC state by incubation at 5˚C. Lanes 1, 5, 9, 11, and 12 contain a 123 bp ladder. Lanes 2-4 contain cells entering the VBNC state at 0, 1, and 2 days; lanes 6-8 contain cells entering the VBNC state at 3, 4, and 5 days; lane 10 contains cells entering the VBNC state at 7 days. Reprinted with permission from: Warner JM, Oliver JD. Appl Environ Microbiol 1998; 64:3025-3028.

Fig. 1.8. (opposite) Effect of plasmid FAC510 carriage in P. fluorescens on culturability in river water at different temperatures. The same P. fluorescens strain was without the plasmid (, “parent”), with the plasmid (❍, “plasmid”), or with the entire plasmid inserted into the chromosome (, “chromosome”). Cells incubated at 5˚C (A) or at 37˚C (B). Plate counts were performed on L agar. Reprinted with permission from: Oliver JD, McDougald D, Barrett T et al. FEMS Microbiol Ecol 1995; 17:229-238.


11

12


Fig. 1.9. P. fluorescens placed into soil microcosms at 5˚(), 25˚(❍), and 37˚C (). Shown are plate counts for each temperature. S. Bunker and J.D. Oliver (unpublished).

1.10. Significance of the VBNC State in the Release of Genetically Modified Bacteria There are numerous practical consequences of the existence of the VBNC state in bacteria. One example is the fact that coliforms, the assay for which is employed worldwide as an indicator of fecal contamination of drinking and recreational waters, also enter the VBNC state. Entrance of these bacteria into this dormant state appears to result from exposure to low temperatures or to increased salt levels. Thus, it must frequently be the case that waters are considered to be “coliform negative” when, in fact, there is a large number of such indicator bacteria present, but in the nonculturable state! Another practical example is that of pathogenic bacteria, many of which are known to enter the VBNC state (see Table 1.1). If oysters are maintained on ice (as they should be!) for several days, this may be sufficient time to induce the VBNC state in V. vulnificus, which typically occurs at high levels in oysters (at least, in the Gulf Coast states of the United States). Public health officials who might examine such oysters for the presence of this species would then conclude the oysters were free of this potentially fatal human pathogen. As we have shown, however, cells of V. vulnificus in the VBNC state retain virulence, and remain capable of producing lethal infections when injected into laboratory mice.22 For a recent review on the public health significance of the viable but nonculturable state of pathogenic bacteria, see Oliver.23 Concern must also be present for researchers and government regulators involved in the release of genetically modified organisms (GMOs). Several bacterial species which are being considered, and even currently employed, for release studies are known to enter the VBNC state (Table 1.1). These include Pseudomonas fluorescens and P. syringae, both of which are induced into the VBNC state by incubation at the high temperatures (ca. 37˚C) which these cells would likely be exposed to in many environments.10 Other factors, such as


13

nutrient deprivation, may also be important for the induction of the VBNC state in pseudomonads. Wilson and Lindow17 have shown that cells of P. syringae which are actively growing on leaves or which were only recently inoculated to leaf surfaces remain fully culturable. However, their study showed that culturable counts of such cells, after 80 hours of epiphytic growth under constant environmental conditions, were two- to four-fold less than direct viable counts, indicating that up to 75% of the population had entered the VBNC state. As these authors stated, “This finding is particularly relevant to the monitoring and detection of GEMS [genetically engineered microorganisms] in terrestrial ecosystems, because if part of an epiphytic population has become metabolically inactive . . . the viable bacterial population size will be underestimated by the plate count method. The plate count may indicate that a GEM has disappeared from the ecosystem, whereas actually, the GEM is still present in low numbers in a viable but nonculturable, quiescent state.” The presence of bacterial cells in the VBNC state presents a major new dilemma in that the ability to monitor the persistence and spread of released GMOs is critical. The routine methods (primarily platings) for detecting bacteria in natural environments can not be employed for cells in the VBNC state. It also appears from our studies that alternate methods to detect such cells, such as PCR amplification, may not be practical alternatives. It must also be asked whether cells in this state are able to either donate or take up genetic material (plasmid or chromosomal DNA) to/from other bacteria which are present as part of the normal microflora. If such gene exchange is possible (although this has not been shown to date), then the problem is compounded, as not only would these GMOs be undetectable, but they might take part in the development of previously non-existent strains of bacteria. Most studies to date on VBNC cells from a variety of genera suggest that plasmids are not lost during entry into this state, suggesting that the acquisition of any new genes would likely take place in an otherwise relatively stable genetic environment. To conclude on a more positive note, our recent studies using certain genetically-tagged bacteria suggest some hope in our ability to detect bacteria present in natural environments in the VBNC state. The “green fluorescent protein” (Chapter 7) is an extremely stable protein which is coded for by the gfp gene present in certain marine jellyfish, and which fluoresces when irradiated with ultraviolet light. The use of this protein is currently finding considerable interest as a marker, following insertion of the gene into the DNA of bacteria intended for environmental release. In collaboration with Dr. Janet Jansson and researchers in her laboratory at the University of Stockholm, we have found that cells producing this fluorescent protein continue to fluoresce even when induced into the VBNC state. Indeed, as seen in Fig. 1.10, not only does a gfp-labeled strain of P. fluorescens continue to fluoresce as it becomes nonculturable, but it appears to fluoresce brighter than logarithmic phase (actively metabolizing) cells of the same gfp-labeled strain. If these results are corroborated in the field, then the use of this marker for monitoring released cells may be of significant value in providing a means of identifying released bacteria even when they are no longer culturable. Such field studies are currently in progress in the author’s laboratory.

1.11. Conclusion Since the pioneering studies from Colwell’s laboratory,24,25 it has become evident that many gram-negative bacteria enter into a state of dormancy in response to one or more of a variety of environmental stresses, including both low and high temperatures. VBNC cells remain metabolically active, although they can not be cultured by standard methods. Cells undergo a variety of morphological, physiological, and biochemical changes as they enter this state, all of which may allow the cells to survive what could otherwise be lethal conditions. On removal of the inducing stress, these cells can resuscitate from the VBNC state, again becoming metabolically active.


14

Fig. 1.10. Fluorescence due to the green fluorescent protein in P. fluorescens, as detected by flow cytometry. Shown is the percent of control cell fluorescence of cells incubated at 5˚C, 30˚C, and 37.5˚C for up to 19d. Cells incubated at 37.5˚C were completely nonculturable by day 15 in this study. M. Lowder and J.D. Oliver (unpublished).

Unfortunately, it appears that cells in the VBNC state may undergo changes in their chromosomal DNA which prevents their detection through PCR amplification. Further, it is possible that VBNC cells may participate in genetic exchange with bacteria making up the normal microflora of a release environment. Such an event could involve gain or loss of plasmids, an event having potentially dramatic consequences on the conditions inducing the VBNC state. Whether these dormant cells can be genetically transformed or transduced is not know at this time, but if so, such events could lead to significant genetic modifications in these nonculturable cells. Thus, the VBNC state presents a number of potential concerns to the researcher and regulator alike, making detection of the introduced cells difficult, and presenting a situation wherein gene exchange could potentially occur, yet be undetectable. Whether or not new technologies such as the use of the gfp marker gene will at least provide a means for monitoring such cells is currently under investigation.

Acknowledgments J.D. Oliver is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Oliver JD. Formation of viable but nonculturable cells. In: Kjelleberg S, ed. Starvation in Bacteria. New York: Plenum Press 1993: 239-272. 2. Kogure K, Simidu U, Taga N. A tentative direct microscopic method for counting living marine bacteria. Can J Microbiol 1979; 25:415-420. 3. Duncan S, Glover LA, Killham K et al. Luminescence-based detection of activity of starved and viable but nonculturable bacteria. Appl Environ Microbiol 1994; 60:1308-1316.


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4. Rattray EAS, Prosser JI, Killham K et al. Luminescence-based nonextractive techniques for in situ detection of Escherichia coli in soil. Appl Environ Microbiol 1990; 56:3368-3374. 5. Kaprelyants AS, Kell DB. Rapid assessment of bacterial viability and vitality using rhodamine 123 and flow cytometry. J Appl Bacteriol 1991; 72:410-422. 6. Paszko-Kolva C, Shahamat M, Yamamoto H et al. Survival of Legionella pneumophila in the aquatic environment. Microbiol Ecol 1991; 22:75-83. 7. McGovern VP, Oliver JD. Induction of cold responsive proteins in Vibrio vulnificus. J Bacteriol 1995; 177:4131-4133. 8. Linder K, Oliver JD. Membrane fatty acid and virulence changes in the viable but nonculturable state of Vibrio vulnificus. Appl Environ Microbiol 1989; 55:2837-2842. 9. Wolf PW, Oliver JD. Temperature effects on the viable but nonculturable state of Vibrio vulnificus. FEMS Microbiol Eco. 1992; 101:33-39. 10. Oliver JD, McDougald D, Barrett T et al. Effect of temperature and plasmid carriage on nonculturability in organisms targeted for release. FEMS Microbiol Ecol 1995; 17:229-238. 11. Oliver, JD, Nilsson L, Kjelleberg S. Formation of nonculturable cells of Vibrio vulnificus and its relationship to the starvation state. Appl Environ Microbiol 1991; 57:2640-2644. 12. Oliver JD. The viable but nonculturable state in the human pathogen Vibrio vulnificus. FEMS Microbiol Lett 1995; 133:203-208. 13. Nilsson L, Oliver JD, Kjelleberg S. Resuscitation of Vibrio vulnificus from the viable but nonculturable state. J Bacteriol 1991; 173:5054-5059. 14. Steinert M, Emody L, Amann R et al. Resuscitation of viable but nonculturable Legionella pneumophila Philadelphia JR32 by Acanthamoeba castellanii. Appl Environ Microbiol 1997; 63:2047-2053. 15. Whitesides MD, Oliver JD. Resuscitation of Vibrio vulnificus from the viable but nonculturable state. Appl Environ Microbiol 1997; 63:1002-1005. 16. Oliver JD, Hite MF, McDougald, D et al. Entry into, and resuscitation from, the viable but nonculturable state by Vibrio vulnificus in an estuarine environment. Appl Environ Microbiol 1995; 61:2624-2630. 17. Wilson M, Lindow SE. Relationship of total viable and culturable cells in epiphytic populations of Pseudomonas syringae. Appl Environ Microbiol 1992; 58:3908-3913. 18. Brauns LA, Hudson MC, Oliver JD. Use of the polymerase chain reaction in detection of culturable and nonculturable Vibrio vulnificus cells. Appl Environ Microbiol 1991; 57:2651-2655. 19. Warner JM, Oliver JD. Randomly amplified polymorphic DNA analysis of starved and viable but nonculturable Vibrio vulnificus cells. Appl Environ Microbiol 1998; 64:3025-3028. 20. Bej AK, Harold N, Vickery MCL et al. Use of PCR to determine genomic diversity and distribution of siderophore-mediated iron acquisition genes in clinical and environmental isolates of Vibrio vulnificus. Abstr Annu Meet Amer Soc Microbiol 1997; Q177, p.485. 21. Mickey S, Warner JM, Simpson LM et al. Detection of enterohemorrhagic Escherichia coli in the viable but nonculturable state using a RAPD-PCR protocol. Abstr Annu Meet Amer Soc Microbiol 1997; P78, 449. 22. Oliver JD, Bockian R. In vivo resuscitation, and virulence towards mice, of viable but nonculturable cells of Vibrio vulnificus. Appl Environ Microbiol 1995; 61:2620-2623. 23. Oliver JD. Public health significance of viable but nonculturable bacteria. In: Colwell RR, Grimes J eds. Nonculturable Organisms in the Environment. ASM Press, 1999 (in press). 24. Roszak DB, Colwell RR. Survival strategies of bacteria in the natural environment. Microbiol Rev 1987; 51:365-379 25. Xu HS, Roberts NC, Adams LB et al. Survival and viability of nonculturable Escherichia coli and Vibrio cholerae in the estuarine and marine environment. Microb Ecol 1984; 8:313-323.

CHAPTER 2

The Use of Antibiotic Resistance Gene Markers for Studying Bacterial Populations in Natural Environments Sharon Egan and Elizabeth M H Wellington

2.1. Introduction

A

ntibiotic resistance genes have been used to mark bacteria by providing a readily selectable phenotype, which can be detected using selective growth media. Detection and monitoring is therefore culture-dependent. A wide range of resistance genes have been characterised (Tables 2.1 and 2.2) which confer resistance to commercially available, inexpensive antibiotics. In addition, resistance genes have provided a valuable tool for cloning and genetic manipulation procedures in both prokaryotes and eukaryotes. Many industrial strains carry resistance genes which have resulted from cloning procedures, enabling selective maintenance of a construct or plasmid carrying the resistance gene by inclusion of the respective antibiotic in the growth medium (For details see ref. 1). The main environmental application for antibiotic resistance gene markers has been the monitoring of gene transfer associated with plasmid or transposon mobility. In addition, the tracking of bacterial populations in soil, water or other environments, and determination of the extent of plant root or shoot colonisation has relied on antibiotic resistance phenotypes.2-4 A distinction can be made between the use of antibiotic resistant mutants and the introduction of cloned resistance genes. The former have been used in field releases where it was necessary to avoid releasing antibiotic resistance genes into the environment.2 The selection provided by antibiotics has provided a powerful tool for isolation of marked bacteria and cultivation based assays have allowed quantitative estimates of cell numbers. Resistance genes have also been detected using molecular methods such as PCR, and this approach provides confirmation of the phenotype. Due to ergotropic and clinical use, some resistance genes have become widely disseminated in the environment and are less useful as selective markers. The choice of resistance gene is therefore important to avoid certain common genotypes. For field releases ethical considerations are important to avoid the use of any resistance gene which may become disseminated to the indigenous bacterial community and thus ultimately contribute to the clinical problems of antibiotic resistant pathogens. The aim of this chapter is to provide information useful for the selection and exploitation of resistance genes as markers for detection and enumeration of bacteria in natural environments.

Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.


18

Table 2.1. Some examples of common bacterial resistance genes and their mode of action Gene

Resistance phenotype

Host

Mechanism of resistance

aph(3)-Iva nptII aphII tsr strA ant(2)-I ant(6)-Ia ant(9)-Ia aac(3)-I aac(6)-Ih aac(3)-Ia mgt tetA/B/C tet(M) ampC otrA/B

kanamycin, neomycin kanamycin, neomycin neomycin thiostrepton streptomycin gentamicin, sisomicin streptomycin spectinomycin gentamicin sisomicin, tobramycin gentamicin, sisomicin oleandomycin tetracycline tetracycline β-lactams oxytetracycline

Bacillus circulans Enterobacter Streptomyces lividans S. lividans Enterobacter Pseudomonas E. faecalis Staphylococcus aureus Pseudomonas Acinetobacter spp. E. coli Streptomyces antibioticus Enterobacter Streptococcus pneumoniae gram-negative bacteria Streptomyces rimosus

Phosphorylation Phosphorylation Phosphorylation Target modification Phosphorylation Nucleotidyltransferase Adenylylation Adenylylation Acetylation Acetylation Acetylation Glycosyltransferase Efflux Ribosomal modification Hydrolysis Ribosomal modification/efflux

2.2. Cultivation Based Assays The traditional approach for monitoring bacteria using antibiotic resistance is selective agar plate counting, although the most probable number technique (MPN) can be used for selective broth culture. Agar antibiotic gradient plates, in which the antibiotic concentration increases linearly from zero to an established level, are produced by making an agar gradient containing the antibiotic and covering a basal agar layer without antibiotic. Such gradients are difficult to replicate and have not generally been used for isolation and counting although they are useful for characterisation of isolates.5 Many antibiotics are unstable, and biological activity can therefore decrease with time or due to inappropriate handling. For example, tetracycline is light sensitive and β-lactams and aminoglycosides, such as neomycin are heat sensitive. Some antibiotics, such as thiostrepton, are only soluble in organic solvents and so will be insoluble in agar forming a suspension, thus affecting availability and activity.6 The bioactivity of antibiotics in agar is affected by many factors (see Hewitt7) and it is well known that they are often complexed by particular constituents of media. Philips8 demonstrated that viomycin exerted a greater selective effect in a defined medium than in a complex one. Methods manuals may often indicate two concentrations of antibiotics, for use in either defined or complex media (For example see ref. 9). The degree of resistance may also be affected by the levels of salt in the cultivation medium; lower salt concentrations can make some bacteria more sensitive.9 Problems can also occur with very polar compounds, for example, streptomycin is a highly water-soluble polar antibiotic and the generation of agar gradients with this compound is particularly difficult as it diffuses very rapidly throughout the agar plate.8 Cultivation-based assays are quantitative, but this procedure is dependent on the bacteria being culturable. The physiological state of the marked bacterium is important and can determine how readily target bacteria can be isolated on highly selective media. The stress of inoculation into a nutrient poor environment such as soil may result in the inoculant becoming nonculturable on isolation media (see Chapter 1) or becoming susceptible

Antibiotic Resistance Gene Markers

19

Table 2.2. Antibiotic resistance genes used as markers Resistance phenotype

Gene/ transposon

Host

Detection1

Reference

kanamycin

aph/ nptII

Enterobacter agglomerans

SP, PCR

Selenska et al14

Escherichia coli

SP, MPN-PCR

Recorbet et al15

kanamycin

nptII

kanamycin

nptII (Tn501) E.coli

SP, probing

Zeph and Stotzky33

kanamycin

nptII (Tn903) Erwinia carotovora

SP

Orvos et al34

kanamycin

nptII (Tn5)

Azospirillum sp.

SP, MPN probing

Bentjen et al35

kanamycin

nptII (Tn5)

P.fluorescens

SP, MPN-PCR

van Overbeek et al32

neomycin

nptII (Tn5)

R.leguminosarum

SP, probing

Hirsch and Stokes2

neomycin

nptII (Tn5)

Rhizobia

SP, probing

Armager and Delgutte44

erythromycin

erm

Bacillus

SP

Wendt/Potthof et al55

kanamycin

nptII

P. fluorescens

SP

Smit and van Elsas24

gentamicin

aadB

chloramphenicol cam

B. subtilis

SP

McDonald et al46

kanamycin

Tn5

E.agglomerans /E.coli

SP

Klingmüller50

neomycin, thiostrepton viomycin

aphII tsr vph

Streptomyces lividans SP

Herron et al6

thiostrepton

tsr

S. lividans

SP, probing

Marsh and Wellington31

tetracycline

unknown

B. subtilis

SP

Amner et al21

neomycin

aphII

Streptomyces violaceolatus/ S. lividans

SP

Wellington et al16

thiostrepton viomycin

tsr vph

1SP: Selective plating, MPN-PCR: Most-probable number-PCR.

20


to selective components in the medium such as antibiotics. Under nutrient limitation some bacteria have been demonstrated to be less susceptible to antibiotics.10,11

2.3. Direct Molecular Monitoring Methods Since the development of the polymerase chain reaction (PCR), molecular monitoring methods for resistance genes have proved to be sufficiently sensitive for detection of sparse populations in natural environments and now provide a valuable adjunct to cultivation assays. However, viability and cellular location of a DNA target cannot be established by PCR detection. In addition, problems still exist with quantitation of PCR although a number of methods have been described for quantitative PCR with targets in natural samples (see Chapter 3).12,13 PCR has been used to examine the persistence of genes introduced to environments such as soil, and in the case of resistance genes, the marker has allowed comparison of molecular and cultivation based detection. Selenska et al 14 targeted the chromosomally-located resistance gene markers aph and nptII. PCR amplification of these genes from total community DNA allowed detection of these genes 56 days after inoculation, even though the Enterobacter agglomerans kanamycin-resistant host could no longer be detected on agar plates. This discrepancy indicated that either the DNA was extracellular and had been released from dead bacteria, or that cells were not culturable. Recorbet et al15 determined the survival of an Escherichia coli population in soil using thenptII gene, and noted that the population decreased to < 102 after 15 days, as calculated by selective plate counts. However, by use of the most-probable-number-PCR method, 5 x 105 target sequences could still be detected after 40 days. It was not clear if this apparent population remaining in the soil was viable but uncultured or simply represented dead cells and extracellular DNA.

2.4. Choice of Antibiotic Resistance Gene Markers The choice of resistance gene markers depends on the environmental use of the marked bacterium and is influenced by the background resistance of the indigenous bacteria in a given habitat. Members of some genera, including Pseudomonas, Mycobacterium and Rhodococcus, have nonspecific mechanisms for resistance to antibiotics and heavy metals, including exclusion from the cell.16 The levels of resistant indigenous bacteria depends on the antibiotic and can range from 103-104 cfu g-1 dry soil for tetracycline and ampicillin resistance17 to 108 cfu g-1 for tobramycin resistance.18 Amner et al19 suggested that examination of the antibiotic resistance profiles of indigenous populations and resistance genes within the gene pool of microbial populations from different environments could be useful in the development of marker systems. The fate of thermophilic actinomycetes in compost was monitored,20 where all microbial constituents of the microflora were sensitive to tetracycline. This resistance gene was thus suitable for introduction into host vector systems to allow the detection of host organisms.21 A plasmid encoding tetracycline resistance was used to monitor Bacillus subtilis in compost. The use of just a single resistance marker was feasible due to the low level (103) of tetracycline resistant isolates in the indigenous bacteria. 21 Conversely, Smalla et al22 examined the distribution of kanamycin resistance in bacteria from different habitats and demonstrated that the resistance phenotype was widely distributed among indigenous bacteria in most habitats examined. Multiple-resistance phenotypes may overcome the problems of background resistance in indigenous bacteria. The use of combined markers for phenotypes which occur less frequently in bacteria, such as viomycin and thiostrepton resistance, is successful and can increase selectivity for an introduced strain. Certain antibiotic resistances often have a low occurrence in indigenous bacteria because of the rarity of use of the antibiotic in medicine,


21

veterinary science or agriculture. In addition, selection from antibiotic-producing bacteria must also have been absent. The combination of two resistance gene markers was used by Cresswell23 to track plasmid-containing streptomycetes in soil. The tsr and nptII genes for thiostrepton and neomycin resistance, respectively, were rarely found in actinomycetes, which also allowed monitoring and detection by PCR. A study by Smit and van Elsas24 used a combination of kanamycin and gentamicin resistance which lowered the level of resistant indigenous bacteria from 105-104 cfu g-1 of dry soil for kanamycin to 103 isolates resistant to both kanamycin and gentamicin. A marker gene cassette with nptII (kanamycin resistance) and aadB (gentamicin resistance) was constructed because these two genes were considered to have a low probability of phenotype and/or genotype prevalence in the indigenous soil populations. This gene cassette was cloned into both a broad-host-range mobilisable plasmid and into a disarmed transposon which would allow tagging of the host’s chromosome. Alternatively, additional markers can be incorporated into hosts, which allow differentiation between the inoculated host and background indigenous bacteria. Van Elsas et al25 engineered an additional unexpressed gene, pat, into the RP4 plasmid of Pseudomonas fluorescens which allowed specific identification of the introduced population by selective plating using the resistance of RP4, followed by DNA hybridisation. Similarly, McNaughton et al26 combined the use of two markers, kanamycin resistance and xylE, on a plasmid introduced into soil. Kanamycin was used in the selective isolation of resistant bacteria, including the inoculant population. Isolates were then examined for catechol oxidation to unequivocally identify the inoculant. When multiple resistance markers have been combined with additional markers for definitive identification, they have proved useful in initial selective procedures.

2.5. Cross-Resistance Resistance to the same antibiotic can be achieved by different genes coding for enzymes with various modes of action or the same gene can confer resistance to more than one antibiotic (Table 2.2). Resistance to paromomycin is conferred by either a phosphotransferase or an acetyltransferase.27 Other resistance gene products can confer a ‘cross-resistance’ to different antibiotics, for example, Skeggs et al28 have shown that an RNA methylase aminoglycoside resistance determinant can confer resistance to kanamycin, apramycin and gentamicin. The aac resistance gene which encodes an acetyltransferase in Streptomyces fradiae and the phosphotransferase-encoding nptII from Tn5 confer resistance to both neomycin and kanamycin.9,29 This functional diversity in resistance genes means that the same phenotype can be due to quite different genes. Unequivocal identification is only possible with selective isolation followed by a molecular method to confirm the presence of the specific resistance marker.17,30-32

2.6. Marking of Bacteria with Antibiotic Resistance Genes Chromosomes of bacteria are often marked by the use of random or site-specific integration vectors, such as transposons. Location of marker genes on the chromosome instead of plasmids may improve stability, and although this reduces the likelihood of a transfer event, it does not eliminate it. Insertion of markers into the chromosome has been achieved by random integration of transposons which encode resistance genes, such as Tn501 in E. coli,33 Tn903 in Erwinia carotovora,34 Tn5 in Azospirillum35 or Pseudomonas fluorescens.32 The fate of bacteria introduced into different environments has been monitored using these transposons. One of the best characterised systems is that for monitoring Rhizobium strains in soil where markers have been developed to monitor numerous field releases of engineered rhizobial strains. Rhizobium leguminosarum was marked by the insertion of Tn5,2 which allowed selection

22


for neomycin resistance on isolation media. This phenotype was not found in native rhizobia, which made it a suitable marker, particularly in combination with the spontaneous chromosomally located rifampicin resistance mutation. Antibiotic resistance gene markers have been inserted on multiple and single copy plasmids for monitoring the fate of inoculants and gene transfer in natural environments. The choice of chromosomal or plasmid location is dependent on the requirements of the study, but gene dose effects with multicopy plasmids will greatly improve molecular detection of both DNA and mRNA. In addition, enhanced phenotypic resistance from multiple copies of the resistance gene marker will aid selection.9 Both naturally occurring R-plasmids and constructed plasmids with one or multiple resistance genes are readily available. However, R-plasmids are mainly found in gram-negative hosts and many will not be able to replicate in gram positive backgrounds. Plasmids from gram-negative and gram positive bacteria have very different conjugation systems and genes involved in plasmid transfer, replication and maintenance are very diverse.36 For gram positive bacteria, resistance plasmids have been constructed (see next section 2.7). For example, Wipat et al 37 used tsr to mark a multicopy streptomycete plasmid which was then introduced into Streptomyces lividans via transformation of protoplasts. This allowed the monitoring of survival and spread of S. lividans in soil microcosms.37 Many resistance marker genes can function in different bacterial species; for example, the nptII gene has been used in many different genera (Table 2.2), but other marker systems are restricted to specific groups. The host range of the vector must also be considered. Although the nptII gene occurs frequently in diverse bacteria, it is often associated with different transposons (Table 2.2). The choice of promoters for marker genes is also an important consideration. The nptII gene from Tn5 is expressed in Streptomyces, but the level of kanamycin resistance varies from 2-200 mg/ml, depending on promoter strength and copy number of the plasmid.9

2.7. Use of Antibiotic Resistance Genes to Monitor Gene Transfer in Soil Antibiotic resistance markers have been used to study gene mobility and dissemination of antibiotic resistance genes has been the subject of a large number of studies due to the problem of drug resistance in bacterial pathogens. Resistance genes are highly mobile under intense selection pressure in clinical environments and have been found associated with chromosomes, plasmids, integrons and bacteriophages.38,39 The application of antibiotics such as streptomycin to treat bacterial rots in soft fruit has resulted in the spread of selected streptomycin resistance genes in Pseudomonas species.40 In addition, genes involved in streptomycin production by streptomycetes are more widely distributed than was previously thought.41,42 There is evidence for the actual presence of antibiotics in soil. Phenazine43 and thiostrepton16 have been detected in soils following the introduction of the respective producer strain. Such antibiotics produced in situ could provide selection pressure for resistance in nature, but at a lower level compared with agricultural, veterinary and clinical environments. Overall, the extent of selection in the natural environment is not well defined. Hirsch and Spokes2 used the Tn5 transposon carrying the nptII gene to mark Rhizobium leguminosarum. and introduced 1012 culturable cells of the strain into soil. Survival was monitored by plating onto selective agar, followed by hybridisation with a Tn5 probe. This study demonstrated that no transconjugants were found, although it was suggested that transfer could have occurred below the limit of detection (< 0.02% isolates received Tn5).2 The inoculant strain persisted in the soil and could be detected seven years later, even in soils in which non-cereal crops were cultivated. A similar study by Armager and Delgutte44 examined the transfer of a Tn5 tagged conjugative plasmid in the same Rhizobium strain in


23

the rhizosphere of different plants and also failed to detect transfer of Tn5 to other rhizobia and even to introduced potential recipients. Other antibiotic resistance gene markers have been used to study plasmid transfer in the natural environment and to examine the extent of conjugation in soil. Cresswell monitored the transfer of plasmid pIJ673 containing neomycin (aphII), viomycin (vph) and thiostrepton (tsr) resistance genes. The plasmid was a derivative of pIJ101.23 Putative transconjugants were selected by growth on neomycin plus thiostrepton-containing agar plates and plasmid presence confirmed by colony hybridisation with a labeled plasmid as a probe. This technique was used to show inter-specific plasmid transfer in Streptomyces strains in both sterile and non-sterile soil batch microcosms.6,30 High copy number plasmids also provide an advantage by effectively increasing the copy number of the gene, which can enhance detection by molecular methods. In a study by Smit and van Elsas24 a plasmid and a chromosomally-inserted transposon were stably maintained in Pseudomonas fluorescens R2f in soil for 7 days and a broad host range plasmid (IncQ) could be mobilised into several indigenous bacterial species. McDonald et al45 examined the persistence of a plasmid encoding chloramphenicol resistance in B. subtilis and reported that plasmid stability in the inoculant population remained at 100% after 28 days. Other studies have shown that engineered plasmids are not stably maintained by the hosts, particularly when released into the environment.46,47 It could be interpreted that in these cases the burden of maintaining multicopy plasmids is greater than the advantage conferred on the host by the marker gene phenotype.48 Examination of intrageneric plasmid transfer in soil has shown that Tn5 labeled plasmids in both E. agglomerans and E. coli could transfer to homologous strains in soil, and the transfer frequency was enhanced by the input of particular nutrients into the system.49 Herron et al6 used pIJ673 (containing nptII, vph and tsr) to monitor the selective effect of antibiotics on the plasmid transfer and survival of Streptomyces in soil microcosms. This study demonstrated that addition of neomycin and thiostrepton to soil microcosms increased the levels of resistant transconjugants from 103 to 105. The population dynamics of bacteriophage and their hosts in soil has also been studied by marking phage with antibiotic resistance genes. Marsh and Wellington31 examined the lysogenic infection of indigenous soil Streptomyces species by using a bacteriophage containing a thiostrepton resistance gene to mark bacteriophage and then select for host resistance in soil microcosms. The addition of tsr marked KC301 actinophage lysates to nonsterile soil gave rise to thiostrepton resistant indigenous streptomycetes which were subsequently shown to contain KC301 DNA by colony blots. Probing digested chromosomal DNA of some of the lysogens showed that their chromosomes contained KC301 prophage.

2.8. Ethical Concerns Although there are obvious advantages in the use of antibiotic resistance gene markers, their value for field releases has been limited. This is probably due to the ethical questions raised about deliberately releasing resistance genes into the environment, which presents the risk of spread into the indigenous bacterial population. However, transfer is known to be limited in conditions of low nutrients24,16 and is also affected by the activity and spatial distribution of bacteria in addition to physical conditions in the soil.50 The problem of transfer has been addressed with the development of modified vectors which inhibit the movement of resistance genes into new hosts.51,52 There is widespread resistance in indigenous bacteria to selected antibiotics in current use in agriculture such as streptomycin. However, there will soon be restrictions on the use of antibiotics in animal growth promotion to reduce the future development of resistance in enteric bacteria. There is a clear link between the development of resistance in antibiotic-fed farm animals and the dissemination

24


of antibiotic resistance genes in enteric bacteria in the environment and within pathogenic groups.53 These data demonstrate the horizontal transfer of genes in natural populations and their selection under conditions of antibiotic administration. Therefore, selection pressure is the key problem. Certain antibiotic resistance gene markers have been proposed for environmental use due to the lack of clinical application of the antibiotic used for selection, thus reducing the chance of resistance development. However, it is often uncertain which chemical classes of antibiotic will prove to be clinically useful in the future and how far cross resistance will affect the use of chemically similar antibiotics. In the case of the antibiotic avoparcin, a glycopeptide fed to pigs and poultry, there was not sufficient difference compared to vancomycin, the clinically important glycopeptide, to prevent cross resistance developing.53 As already discussed in Section 2.6, some field releases have been approved which used antibiotic resistance gene markers for tracking genetically modified micro-organisms (GMMs) and included Tn5 and the kanamycin resistance gene aph(3')-II (see Chapter 8). The risk of a gene transferring from GMMs into the environmental gene pool is presumably greater than for a transgenic crop plant marked with an antibiotic resistance gene. The only GMM-based product containing an antibiotic resistance gene for marketing within the European Union is a kit consisting of Streptococcus thermophilus modified with the cat (chloramphenicol acetyltransferase) gene. The cat gene is located on a plasmid, pMJ723, harbouring the lux genes, and detects antibiotic residues in milk. The strain is not released into to the environment, as it is contained in vials throughout the test. For transgenic crop plants, the marker antibiotic resistance genes aph(3')-II and blaTEM-1, for kanamycin/neomycin resistance and β-lactam resistance respectively, have been approved for marketing within Europe. Future research should address the levels of transfer of these genes to indigenous micro-organisms, as plant DNA containing aph(3')-II is known to successfully transform suitable, competent, recipient bacteria. 54

2.9. Conclusion Antibiotic resistance genes have a long history of reliability for use as highly selective, versatile markers allowing detection and enumeration of bacteria in environmental samples. Resistance genes found in non-antibiotic producing bacteria are readily expressed in a wide range of host backgrounds. Examples include the neomycin resistance gene, nptII, which has been used in gram-positive (high and low GC) and gram-negative bacteria, yeasts, plant and animal cell cultures. The resistance genes are highly mobile between bacterial groups and this may be a reason for their versatility and often near universal expression. A wide range of genes have been cloned and frequently used with their own promoter in diverse hosts. Levels of expression and strength of resistance may vary slightly from strain to strain but this is also related to the nature of the construct used. Multiple copies of a resistance gene secure high levels of resistance but only under defined conditions of growth and assay. No other type of marker provides such an extensive range of genes for use with almost any bacterium, resulting in a highly selective marker. Some bacteria now carry an extensive range of antibiotic resistance genes resulting from the excessive use of antibiotics for over 50 years and this may reduce the choice of genes useful for marking a strain within a population of close relatives. The background resistance of indigenous bacterial populations may also be a problem where certain phenotypes, such as kanamycin resistance, are widespread in soil bacteria due to the use of antibiotics in agriculture. This problem can be overcome by combining unique sets of resistance genes or by combining resistance genes with markers such as lux or gfp (see Chapters 5-7).


25

Acknowledgments E.M.H. Wellington is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Sambrook J, Fritsch EF, Maniatis T. Molecular cloning: a laboratory manual, 2nd ed. NY: Cold Spring Harbour Laboratory Press, 1989. 2. Hirsch P, Spokes JD. Survival and dispersion of genetically modified rhizobia in the field and genetic interactions with native strains. FEMS Microbiol Ecol 1994; 15:147-160. 3. Kinkel LL, Wilson M, Lindow SE. Utility of microcosm studies for predicting phylloplane bacterium population sizes in the field. Appl Environ Microbiol 1996; 62:3413-3423. 4. Nielsen KM, Bones AM, van Elsas JD. Induced natural transformation of Acinetobacter calcoaceticus in soil microcosms. Appl Environ Microbiol 1997; 63:3972-3977. 5. Philips L, Wellington EMH. The distribution of DNA sequences hybridizing with antibiotic production and resistance gene probes within type strains and wild isolates of Streptomyces species. J Antibiotics 1992; 45:1481-1491. 6. Herron PR, Toth IK, Heilig GHJ et al. Selective effect of antibiotics on survival and gene transfer of streptomycetes in soil. Soil Biol Biochem 1998; 30:673-677. 7. Hewitt W. Microbiological Assays. London: Academic Press, 1977:284-317. 8. Philips L. PhD thesis. University of Warwick, U.K. 1992. 9. Hopwood DA, Bibb MJ, Chater KF et al. Genetic Manipulation of Streptomyces: A laboratory manual. Norwich: The John Innes Foundation, 1985. 10. Oliver JD. The viable but non-culturable state in the human pathogen Vibrio vulnificus. FEMS Microbiol Lett 1995; 133:203-208. 11. Turpin P, Dhir VK, Maycroft K et al. The effect of Streptomyces species on the survival of Salmonella in soil. FEMS Microbiol Ecol 1992; 101:271-280. 12. Picard C, Ponsonnet C, Paget C et al. Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction. Appl Environ Microbiol 1992; 58:2717-2722. 13. Romanowski G, Lorenz M, Wackernagel W. Use of PCR to monitor the persistence of extracellular plasmid DNA introduced into soils. Appl Environ Microbiol 1993; 59:3438-3446. 14. Selenska S, Schenzinger S, Klingmuller W. Direct detection of particular DNA sequences in soil. In: Gauthier MJ, ed. Gene transfers and environments. Proceedings of the third European meeting on Bacterial Genetics and Ecology (BAGECO-3). Springer-Verlag 1992:3-8. 15. Recorbet G, Picard C, Normand P et al. Kinetics of the persistence of chromosomal DNA from genetically engineered Escherichia coli introduced into soil. Appl Environ Microbiol 1993; 59:4289-4294. 16. Wellington EMH, Herron PR, Cresswell N. Gene transfer in terrestrial environments and the survival of bacterial inoculants in soil. In: Edwards C, ed. Monitoring genetically Manipulated Microorganisms in the Environment. Chichester: John Wiley and Sons. 1993: 137-170. 17. van Elsas JD, Trevors JT. Plasmid transfer to indigenous bacteria in soil and rhizosphere: problems and perspectives. In: Fry JC and Day MJ eds. Bacterial Genetics in Natural Environments. Chapman and Hall, 1993:188-199. 18. Henschke RB, Schmidt FRJ. Use of wide host range promoters to monitor the fate of recombinant DNA in soil. In: Fry JC and Day MJ eds. Bacterial Genetics in Natural Environments. Chapman and Hall, 1993:200-206. 19. Amner W, Edwards C, McCarthy AJ. Composting as a model system for monitoring the fate of genetically manipulated gram-positive bacteria. In: Edwards C, ed. Monitoring Genetically Manipulated Microorganisms in the Environment. Chichester: John Wiley and Sons, 1993: 83-109.

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20. Amner W, McCarthy AJ, Edwards C. Quantitative assessment of factors affecting the recovery of indigenous and released thermophilic bacteria from compost. Appl Environ Microbiol 1988; 54:3107-3112. 21. Amner W, McCarthy AJ, Edwards C. Survival of a plasmid-bearing strain of Bacillus subtilis introduced into compost. J Gen Mirobiol 1991; 137:1931-1937. 22. Smalla K, van Overbeek LS, Pukall R et al. Prevalence of nptII and Tn5 in kanamycinresistant bacteria from different environments. FEMS Microbiol Ecol 1993; 13:47-58. 23. Cresswell N. PhD thesis. University of Warwick, U.K. 1992. 24. Smit E,van Elsas JD. Conjugal gene transfer in the soil environment; New approaches and developments. In: Gauthier MJ, ed. Gene transfers and environments. Proceedings of the third European meeting on Bacterial Genetics and Ecology (BAGECO-3). Springer-Verlag 1992:79-94. 25. van Elsas JD, Fouchier R, van Overbeek LS. A specific marker, pat, for studying the fate of introduced bacteria and their DNA in soil using a combination of techniques. Plant and Soil. 1991; 138:49-60. 26. McNaughton SJ. Rose DA, O’Donnell AG. Growth and survival of genetically modified Pseudomonas putida in soils of different texture. In: Stewart-Tull DES, Sussman M. eds. The release of genetically modified microorganisms-REGEM 2. New York: Plenum Press,1992, 191-193. 27. Zalacain M, Gonzalez A, Guerrero MC et al. Nucleotide-sequence of the hygromycin-β phosphotransferase gene from Streptomyces hygroscopicus. Nucleic Ac Res 1986; 14: 1565-1581. 28. Skeggs PA, Holmes DJ, Cundliffe E. Cloning of aminoglycoside-resistance determinants from Streptomyces tenebrarius and comparison with related genes from other actinomycetes. J Gen Microbiol. 1987; 133:915-23. 29. Shaw KJ, Rather PN, Hare RS et al. Molecular genetics of aminoglycoside resistance genes and familial relationships of the aminoglycoside-modifying enzymes. Microbiol Rev 1993; 57:138-163. 30. Wellington EMH, Cresswell N, Saunders V. Growth and survival of streptomycete inoculants and extent of plasmid transfer in sterile and non-sterile soil. Appl Environ Microbiol 1990; 56:1413-1419. 31. Marsh P, Wellington EMH. Phage-host interactions in soil. FEMS Microbiol Ecol 1994; 15:99-108. 32. van Overbeek LS, van Veen JA, van Elsas JD. Induced reporter gene activity, enhanced stress resistance and competitive ability of a genetically modified Pseudomonas fluorescens strain released into a plot planted with wheat. Appl Environ Microbiol 1997; 63:1965-1973. 33. Zeph LR, Stotzky G. Use of a biotinylated DNA probe to detect bacteria transduced by bacteriophage P1 in soil. Appl Environ Microbiol 1989; 55:661-665. 34. Orvos DR, Lacy GH, Cairns J. Genetically engineered Erwinia carotovora—Survival, intraspecific competition, and effects upon selected bacterial genera. Appl Environ Microbiol 1990; 56:1689-1694. 35. Bentjen SA, Fredrickson JK, van Voris P. Intact soil-core microcosms for evaluating the fate and ecological impact of the release of genetically engineered microorganisms. Appl Environ Microbiol 1989; 55:198-202. 36. Macrin FL, Archer GL. Conjugation and broad host range plasmids in streptococci and staphylococci. In: Clewell DB, ed. Bacterial Conjugation. London: Plenum Press, 1993: 313-330. 37. Wipat A, Wellington EMH, Saunders VA. Streptomyces marker plasmids for monitoring survival and spread of streptomycetes in soil. Appl Environ Microbiol 1991; 57:3322-3330. 38. Tschäpe H. The spread of plasmids as a function of bacterial adaptability. FEMS Microbiol Ecol 1994;15:23-32. 39. Salyers AA, Shoemaker NB. Broad host range gene transfer: Plasmids and conjugative transposons. FEMS Microbiol Ecol 1994; 15:15-22. 40. Sundin GW, Monks DE, Bender CL. Distribution of the streptomycin-resistance transposon Tn5393 among phylloplane and soil bacteria from managed agricultural habitats. Can J Microbiol 1995; 41:792-799.


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41. Wiener P, Egan S, Huddleston, AS, Wellington EMH. Evidence for transfer of antibiotic resistance genes in soil populations of streptomycetes. Mol Ecol 1998; 7:1205-1216. 42. Egan S, Wiener P, Kallifidas D, Wellington EMH. Transfer of streptomycin biosynthesis gene clusters within streptomycetes isolated from soil. Appl Environ Microbiol 1998; 64:5061-5063. 43. Thomashow L, Weller DM, Bonsall RF et al. Production of the antibiotic phenazine-1carboxylic acid by fluorescent Pseudomonas species in the rhizosphere of wheat. Appl Environ Microbiol 1990; 56:908-912. 44. Amarger N and D. Delgutte. Assesment of gene transfer under field conditions in France. In: Abstracts of: Horizontal Gene transfer- Mechanisms and Implications. Bielefield, Germany. 1994. 45. McDonald IR, Riley PW, Sharp RJ et al. Survival of plasmid-containing Bacillus subtilis released into mushroom compost. Microbial Ecol 1998; 36:51-59. 46. Tokuda Y, Ano T, Shoda M. Survival of Bacillus subtilis Nb22, an antifungal-antibiotic iturin producer, and its transformant in soil-systems. J. Ferment. Bioeng. 1993; 75: 68-73. 47. Crowley DE, Brennerova MV, Irwin C. Rhizosphere effects on biodegradation of 2,5dichlorobenzoate by a bioluninescent strain of root-colonizing Pseudomonas fluorescens. FEMS Microbiol Ecol 1996; 20:79-89. 48. Tang WZ, Pasternak JJ, Glick BR. Persistence in soil of the plant-growth promoting rhizobacterium Pseudomonas putida gr12-2 and genetically manipulated derived strains. Canad J Microbiol 1995; 41 445-451. 49. Klingmuller, W. 1992. Risk assesment in releases of nitrogen-fixing Enterobacter into soil; survival and gene transfer, as influenced by agricultural substrates. In: Stewart-Tull DES, Sussman M, eds. The Release of Genetically Modified Microorganisms-REGEM 2. New York: Plenum Press,1992:155-157. 50. van Elsas JD. Antibiotic resistance gene transfer in the environment: an overview. In: Wellington EMH, van Elsas JD, eds. Genetic Interactions Among Microorganisms in the Natural Environment. Oxford: Pergamon Press, 1992; 17-39. 51. Bej AK, Perlin MH, Atlas RM. Model suicide vector for containment of genetically engineered microorganisms. Appl Environ Microbiol 1988; 54:2472-2477. 52. Sanchis V, Agaisse H, Chafaux J et al. A recombinase-mediated system for elimination of antibiotic resistance gene markers from genetically engineered Bacillus thuringiensis strains. Appl Environ Microbiol 1997; 63:779-784. 53. Witte W. Impact of antibiotic usage in animal feeding on resistance of bacterial pathogens in humans. In: Chadwick DJ, Goode J, eds. Antibiotic resistance: Origins, evolution, selection and spread. Ciba Foundation Symposium 207. Chichester:John Wiley and Sons, 1997 61-71. 54. Gebhardt F, Smalla K. Transformation of Acinetobacter sp. strain BD413 by transgenic sugar beet DNA. Appl Environ Microbiol 1998; 64:1550-1554.

CHAPTER 3

Extraction and Analysis of Microbial Community Nucleic Acids from Environmental Matrices Jan Dirk van Elsas, Kornelia Smalla, Christoph C. Tebbe

3.1. Introduction

E

nvironmental monitoring on the basis of nucleic acids is increasingly being recognized as an extremely powerful approach, since organisms or genes can be directly assessed, even without prior cultivation. Hence, targets present in unculturable, poorly-culturable or as yet uncultured organisms, which would escape detection when using traditional cultivation-based approaches, are assessable. In the last decade, a number of developments, including the rapid development of nucleic acid detection methodologies, have provided a great thrust to the efforts to assess microbes directly in their natural environment. Areas of study that have been significantly stimulated by nucleic acid based approaches are: 1. Monitoring indigenous microorganisms of interest from an ecological or biotechnological perspective, 2. The study of natural microbial diversity and of factors that disturb that diversity, 3. The need to monitor genetically modified microorganisms (GMMs) based on their unique nucleic acid sequences, 4. The assessment of the expression of specific microbial genes in the environment. The environmental matrices to be sampled and extracted may range from a variety of bulk soils, plant roots (rhizospheres and/or rhizoplanes), leaves (phyllosphere) or interior plant tissue, seeds, rockwool and manure to diverse aquatic systems such as sediments and sewages. A large and diverse suite of protocols for the extraction and analysis of nucleic acids from these environments has been collected in recent texts,1,2 and the reader is referred to these for detailed information. The basic principles of these protocols are inherently very similar, as in most cases the nucleic acids are to be obtained from mixed microbial communities that occur adsorbed to, and heterogeneously dispersed in, the matrix under study. In addition, there are often compelling reasons, e.g., when the persistence of a specific target gene has to be monitored, to also analyze the fractions of the total nucleic acid pool that occur extracellularly.3 There are two different approaches to the extraction of nucleic acids from mixed microbial communities in environmental matrices. The first approach is based on the prior separation of microbial cells from the matrix, after which the dislodged cells are lysed and the nucleic acids extracted and further purified (Cell extraction / nucleic acid extraction, or Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.


30

indirect, approach4,5). The second approach is based on the direct lysis of microbial cells in the environmental matrix (e.g., soil), followed by separation of the nucleic acids from the matrix and from cell debris and other impurities (Direct lysis approach6). Major differences between the two strategies are the higher nucleic acid yields obtained with the direct lysis approach, coupled to the co-extraction of larger amounts of contaminating substances7. Moreover, the cell extraction/nucleic acid extraction approach in principle allows the removal of a large fraction of the extracellular nucleic acids from the bacterial cells prior to lysis, thus allowing a better assessment of target DNA present inside the microbial cells in the environment. Since these pioneering studies on DNA extraction from soil,4-7 there has been a considerable methodological development, the main purpose of which was the omission of laborious purification steps, mainly hydroxyapatite column chromatography and cesium chloride (CsCl) gradient purifications, replacing these by faster approaches. Thus, the numerous nucleic acid extraction protocols that are currently in use in different laboratories1-2,8-19 all share a relatively small number of individual extraction and purification steps (Table 3.1).

Table 3.1. Some examples of frequently used steps in nucleic acid extraction and purification protocols Purpose

Step

Principle

Remarks

Cell lysis

Enzymes and detergents

Breakdown and solubilization of cell envelope components

Very diverse requirements of different cells (e.g., grampositives versus gramnegatives.

Freeze/thaw (heat and cold shocks)

Temperature shocks combined with water crystals, to destabilize membranes

Less efficient for lysis of grampositive bacteria

Grinding with liquid N2

Abrasive action of grinding Efficient for fungal with soil combined with spores and ice crystals mycelium

Bead beating using small glass beads and high frequency shaking

Mechanical lysis (brute force)

Recognized as highly efficient in lysing a wide range of bacterial and fungal/yeast cells

Microwave oven

Heat induced lysis

Combined with other lysis methods

Sonication

High energy induced lysis

Often not efficient if energy-dissipating materials are present

Extraction and Analysis of Nucleic Acids

31

Purpose

Step

Principle

Remarks

Extraction and precipitation

Phenol

Denaturing and extractive action on proteins and lipoproteins

Standard method in molecular biology

Chloroform

Denaturing action on proteins and hydrophobes


Ethanol or isopropanol precipitations

Removal of salts and solutes


Polyethylene glycol (PEG) precipitation

Precipitation/concentration of DNA

CsCl precipitation

Removal of impurities by salting-out effect

KAc/NH4Cl precipitations

Precipitation of DNA due to high molarity acetate

Glassmilk sorption

Sorption on glassmilk beads, followed by differential desorption

Highly efficient in removing humic compounds from DNA

Elutip D

Chromatography separation

Method repeated on same extract

Wizard DNA cleanup spin columns

Powerful separation of DNA from humics via chromatography over mini spin column

Sephadex G50/G75/G200

Separation of contaminants, e.g., humics, via gel filtration

Efficient fast method, often combined with other methods

Gel electrophoresis

Charge- and size-related separation of nucleic acids from impurities

Very efficient, but laborious method

PVPP* sorption

Selectively removes humics from DNA/RNA solutions

PVPP needs rigorous acid wash

Hydroxy apatite chromatography

Selective binding of nucleic acids to HAP**, column. Differential elution of DNA or RNA by varying phosphate concentrations

Theoretically very good, but practically difficult. Often low recoveries

Purification

* PVPP: polyvinyl poly pyrrolidine;** HAP: hydroxyapatite

32


Fig. 3.1. Outline of nucleic acid based detection protocols

There have even been efforts to simplify and miniaturize soil DNA extraction protocols to a level where their on site use becomes feasible.20 Most of the current protocols have been shown to produce DNA and/or RNA suitable for the analysis of microbial diversity or microbial fate. However, they often differ in the way they release and lyse microbial cells from the environmental matrix, and such (qualitative and quantitative) differences are likely to affect the final analyses performed. Hence, it is key to our understanding of microorganisms in their natural settings as described by molecular methods, that the possible biases introduced by cell extraction and lysis methods are understood. This chapter will review currently available strategies to recover and purify (1) microbial (bacterial) cells and (2) nucleic acids (DNA and RNA) from environmental matrices, and will then briefly address the use of these nucleic acids in monitoring methods to assess


33

microbial diversity and inoculant fate. Figure 3.1 gives a general outline of these methodologies.

3.2. Extraction of Microbial Cells from Environmental Matrices Recovery and concentration of microbial cells may be an important first step if nucleic acid analysis is to be performed in environments with relatively low cell densities, such as aquatic or phyllosphere habitats. Moreover, even for environments with high population densities, it can be advantageous to prepare a cell fraction prior to nucleic acid extraction, in particular if intracellular targets are to be assessed. Bacterial diversity assessments based on DNA reassociation kinetics21 even depend on the DNA being primarily of bacterial origin. Paul recently reviewed the available methods for cell concentration from aquatic environments;22 a variety of approaches has been used, the most effective being those including the filtration of large volumes through membrane filters.23-25 Tangential flow filtration26 or Vortex flow filtration27 have the advantage that cells experience less mechanical stress and that large volumes (up to 100 L) can be processed within short times. Prefiltration through 3 µm filters can be used to remove large particles which would clog filters.26 Alternatively, high-speed centrifugation is applicable for concentrating bacteria in smaller water volumes. Filters with microbial biomass or bacterial pellets obtained after highspeed centrifugation can be stored frozen until further processing. As most bacterial cells in nature, including aquatic environments, occur attached to surfaces (forming microcolonies or biofilms28), cell dislodgement methods are often required. Cells can actually adhere quite strongly to surfaces by bonding mechanisms such as via bacterial polymers, pili or flagellae, electrostatic forces or water bridging.29-31 Bacteria can also be entrapped in soil aggregates which are formed through gluing by bacteriallyproduced polysaccharides and physico-chemical interactions between silica/clay surfaces and decomposed organic matter.32, 33 The representative extraction of surface-attached bacteria from environmental matrices requires that the cells bound by the various modes and with different strengths are efficiently dislodged. Dissociation of cells from surfaces is generally achieved by repeated homogenization steps. For soil, homogenization can be achieved by shaking soil suspensions with gravel or beads, or by blending in Stomacher or Waring blenders.34 Mild ultrasonication in a low energy bath has also been used.29 However, the amount of energy needed to completely disperse soil aggregates can result in considerable cell death.29,35 During extraction, bacterial cells experience not only mechanical stress but also changes in their physico-chemical conditions. Thus, the physiological state of cells following dislodgment from surfaces, reflected in the diversity of intracellular RNA molecules, might not represent their in situ physiological state. Lindahl et al36 found that the activity of extracted bacterial fractions is often lower than that in soil slurries. Furthermore, Priemé et al31 reported that treatment of soil samples in a Waring blender decreased methane oxidation activity, with a profound difference between different soils. Soil slurries in sodium pyrophosphate showed less than 10% of the methane oxidation observed with soils resuspended in water. Furthermore, the sodium pyrophosphate used might be problematic in microbiological incubations due to the enhanced supply of phosphorus.33 Following dislodgment, separation of bacterial cells from the particulate matter is usually achieved by low-speed centrifugation, with their subsequent recovery in a pellet by high-speed centrifugation (differential centrifugation37). Recent studies revealed that the centrifugal force applied is highly critical;38 centrifugal forces over 500 xg reduced cell recoveries dramatically. The use of forces < 100 xg was therefore recommended, but at these reduced speeds, recovery rates did not exceed 45% of the total discernable cells.38

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Fig. 3.2. A protocol for the recovery of bacterial cells from soil.

An alternative method to disperse soil particles and dissociate microbial cells from soil, sediment or root parts involves the use of cation exchange resins. Shaking of soil with, for instance, Dowex39 or Chelex-10029,40,41 is used to remove the bivalent and polyvalent cations responsible for electrostatic bonding between like-charged bacterial cells and soil particles. Detergents are also used to overcome adhesive interactions. A comparison of five different treatments (Fig. 3.2) showed that cell extraction could be improved by using Stomacher blending instead of shaking. Consistent results and a fairly rapid treatment were possible with the automated paddle action of the Stomacher. On the other hand, Lindahl and Bakken42 did not find any positive effect of the Chelex-100 treatment on cell extraction efficiency and, thus, recommended simple, threefold repeated, blending with water. Shaking in lowelectrolyte concentrations, e.g. in distilled water, increases the interactive free energy between like-charged soil particles. In a recent comparison of both methods with agricultural soil, the total cfu numbers obtained were found to be comparable between the methods (unpublished). However, the water-based protocol yielded a higher number of different colony types, suggesting that the extraction protocol strongly affects the diversity of bacterial types obtained. In particular for soils with high clay contents, separation of bacterial cells and soil particles is necessary. This can be achieved by flotation in the density media Percoll or


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Nycodenz,31,43 but not with simple centrifugation steps, because clay particles and bacterial cells show similar sedimentation behavior. The efficiency of different cell extraction protocols thus depends on the soil type; hence, different protocols may prove suitable for different soils.43 However, even with the most optimized protocol, complete extraction of all bacterial cells, in particular of cells bound to soil particles, is probably impossible. Thus, for studies on the diversity of microbial communities in soil, it seems reasonable that the cells recovered represent, in their relative abundance, the surface-attached microbial community. As microbial cells may occur both at the outside of soil aggregates and in the interior regions, the extent to which these soil aggregates are disrupted will determine the sites from which the inhabiting bacterial cells are dislodged. To monitor the population dynamics of bacterial inoculants, it is, thus, primordial to understand their localization, as the protocol to be used and the limit of detection are affected by it. Moreover, inoculant cells may preferentially occur at different locations than indigenous microbes, and hence the requirements with respect to extraction are different. The bacterial cell fractions obtained can be used for the extraction of genomic DNA4,5 or for the parallel extraction of both DNA and RNA (Fig. 3.1).44,45 Furthermore, the cell fractions can serve for assessments of cell numbers using fluorescent dyes or immunofluorescence enumeration. In general, nucleic acids recovered from soil/rhizosphere bacterial cell fractions are less contaminated with coextracted humic acids, fulvic acids, polyphenols, polysaccharides, or other plant-derived substances which can hinder molecular analyses, than these recovered directly.7 For polluted soils, separation of bacterial cell fractions may even be an absolute requirement. For instance, DNA extracted directly from a zinc-contaminated soil was not PCR amplifiable even after several purification steps, whereas DNA extracted from the recovered bacterial fraction was amplifiable without any problems (Brim et al, unpublished).

3.3 Cell Disruption The aim of the cell disruption step is to lyse as many target microbial cells as possible, since any subsequent analysis on the basis of nucleic acids is greatly sensitized if this step is optimized. The lysis protocol can be targeted towards a specific microbial group or to the total microbial community. Unfortunately, cell lysis is often the main limiting step in nucleic acid extraction protocols, even of pure cultures.46 Due to the enormous differences in lysability of cells (cf. easily lysed enterobacteriaceae or members of the genus Pseudomonas versus recalcitrant types such as Bacillus spores or mycobacteria), different preferred lysis protocols have been concocted. Moore et al 46 summarized the different available methods for a range of bacteria and fungi, and suggested that each microbial group should be tackled by a particular choice or combination of enzymatic, detergent-based and mechanical methods, the aim clearly being the generation of high nucleic acid yields of optimal quality (molecular size and purity). Cell lysis in the presence of an environmental matrix is different from that without it, and is often more difficult, since compounds from the matrix can affect the action of the lysing agents used, and cells can be in a physiological/morphological state that makes them refractory to lysis.47 For example, the presence of soil particles can inhibit the action of lytic enzymes due to effects on pH, ionic strength or to simple adsorption, but enhance the (mechanical) action of mortar/pestle grinding in the presence of liquid nitrogen, due to their abrasive action.48,49 Moreover, it is possible that certain sites in soil are not reached by lytic enzymes, resulting in a lack of lysis in these specific sites. It is, therefore, likely that lysis efficiencies and possible biases are different for lysis steps applied to pre-separated bacterial fractions as opposed to those of cells in the presence of their natural matrix.

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The strategies used in most protocols to achieve lysis of bacterial cells from environmental matrices, either in its presence or absence, are: 1. Enzymatic removal of cell wall material, 2. Chemical action such as solubilization of the cell membrane with detergents, 3. Physical action such as freezing/thawing, ultrasonication or microwave oven treatment, and 4. Mechanical action such as achieved via e.g., bead beating or mortar/pestle grinding. Often, some of these strategies are combined. Key aspects are described below: 1. A range of enzymes that attack different cell wall components can be applied, such as proteinase K, lysozyme, mutanolysin, lysostaphin, subtilisin or achromopeptidase.46 These enzymes are more effective when combined with EDTA. 2. Commonly used detergents are sodium dodecyl sulfate (SDS) and N-lauroylsarcosine (Sarcosyl). Cetyltrimethyl ammonium bromide (CTAB) is also often used. Nonpolar detergents, such as Triton X, Tween or Nonidet P-40, which provide for milder treatments, are less common. 3. Physical methods have mostly been combined with other (enzymatic or mechanical) methods, as there is doubt whether their single use will result in acceptable levels of lysis.15,20 For instance, the application of freeze/thaw steps, even repeatedly, has been shown to be inferior to bead beating.15,20 Moreover, it is difficult to achieve high lysis rates by the single use of ultrasonication of soil slurries. 4. Of particular interest is the use of the bead beater (Braun cell homogenizer), which is able to lyse even very recalcitrant cellular forms such as Bacillus spores, mycobacteria or fungal conidia.20,50,51 This method was proposed in a pioneering paper on direct extraction of soil DNA7 and is still widely in use. Furthermore, a method in which the abrasive action of mortar/pestle grinding was combined with liquid nitrogen snap-freezing was shown to be efficient in releasing fungal DNA.48,49,52 Table 3.1 lists the most o c mmonly used steps of popular protocols. Combinations of these approaches have also been used. For instance, Picard et al53 have shown that a combination of ultrasonication, microwave oven and enzymatical treatment resulted in a higher degree of cell lysis and thus DNA yield from soil, than any of these steps performed in separate. However, DNA shearing was a severe problem. Problems with all these approaches are related to either the uneven distribution of microbial cells throughout environmental matrices, resulting in their uneven exposure to the action of lysis agents, or to differences in the resistance of different microbial types to lysis. It is therefore prudent to state that complete lysis of cells in environmental matrices is most often impossible to achieve. Hence, the data obtained on the basis of environmental DNA (and/or RNA) should be regarded as descriptive of the populations recovered in a relative sense, and rigorous checks on the nature of the possible bias introduced may be necessary.

3.4. Extraction and Purification of DNA 3.4.1. Extraction Generally, the aim of the extraction steps is to quickly separate the microbial DNA from contaminating compounds such as humic acids, fulvic acids, polyphenols, polysaccharides, enzymes (in particular DNAses) and cellular debris. Protocols for pure culture extractions, although similarly structured,46 often do not deal with compounds that abound in environmental extracts, and these protocols are therefore not a priori suitable. There are a limited number of physico-chemical strategies to quickly remove contaminating compounds, which might eventually be combined into a standardized protocol. However, a wide


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range of protocols for extraction of environmental nucleic acids is currently still in use.1,2 The main strategy is based on organic solvents which interact with the hydrophobic parts of proteins and lipoproteins, resulting in their denaturation and extraction. Using, for instance, phenol—or chloroform—water phase separations, many proteins, lipoproteins and other hydrophobic compounds concentrate in the organic phase, whereas nucleic acids will concentrate in the water phase. Phenol is known to be efficient in denaturing and extracting proteins, whereas chloroform interacts with proteins and polysaccharidic material. Hence, a combination of these two extractants is often employed. Chloroform also removes traces of phenol still present in the aquatic phase. In addition to phenol/chloroform extractions, ethanol or isopropanol precipitations of DNA, followed by washes, are often used to remove salts, SDS and other impurities.54 The solution produced in these initial steps, called the crude extract, will often be a rather impure aqueous solution of nucleic acids. DNA and RNA show similar behavior and may be both present in these crude extracts.

3.4.2. Purification The aim of purification of the crude extract is to recover the nucleic acids from the crude extract in a form ready for analyses by appropriate molecular techniques. Ideally, the procedure is as rapid and simple as possible, and incurs minimal quantitative and/or qualitative loss of, or damage to, the nucleic acids. The different protocols currently in use in different laboratories have been based on an array of different steps in various combinations. Table 3.1 presents a summary of purification methods employed in selected protocols. There is ample evidence that the requirements, with respect to DNA purity, for restriction by different enzymes, cloning, PCR amplification, reverse transcription or direct hybridization can be quite different. Moreover, different enzymes, e.g., thermostable DNA polymerases, have different requirements as to DNA purity.14,55 Therefore, for each new experimental system, the extent to which nucleic acid purification is needed has to be carefully established and controlled. However, a few “trends” in the purification of environmental DNA are worth mentioning. The powerful purification steps used in the early protocols, such as CsCl gradients5 and hydroxyapatite chromatography,4,30 have been found to be either very tedious or unpredictable with respect to yields. Gel electrophoresis has been shown to offer a powerful (one-step) alternative strategy to obtain DNA pure enough for molecular analyses via PCR.18,19,56, 57 Young et al58 even used polyvinylpolypyrrolidone in their gels to enhance the selective removal of humics. However, gel electrophoresis may lead to a bias when the DNA is subsequently used for PCR amplification and community fingerprinting.58 Moreover, the method can be tedious, and has therefore been superseded by other approaches. Thus, there has been an extensive search for reliable alternative purification methods that were simple, fast and provided high throughput.8-12,15-19 Often, these protocols relied on a sequence of short, different, treatments, including e.g., KAc or NH4Ac precipitations,8,11 CTAB extraction,10 hydroxyapatite mini-spincolumns12 and Sephadex gel filtration.8,12,56,59,60 One rapid protocol that consistently yielded PCR amplifiable and restrictable DNA from different soils, originally developed by Smalla et al,15 was based on a two- or three-step purification consisting of sequential KAc precipitation, CsCl precipitation and (originally) glass milk or Wizard spin column purification. In later work, it was shown that this protocol was very flexible and adaptable to a wide range of soil types.61 In a parallel study, Zhou et al 62 also described a protocol which contained four flexible purification routines applicable to a range of different soils. As previously mentioned, a recent development aimed at the on site extraction of nucleic acids from soil. A very simple one-step purification over a Sephadex G-200 column was

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found to yield DNA sufficiently pure for PCR amplification and detection of key microbial groups.20 However, it is doubtful the DNA obtained was of a quality similar to that obtained by more rigorous protocols.

3.5. Analysis of DNA and Detection of Specific Sequences Commonly, the quantity and quality of extracted environmental DNA is analyzed by standard methods, such as absorbance at 260 nm in the spectrophotometer or gel electrophoresis.1,2,54 However, due to the presence of coabsorbing compounds (humic acids) in most nucleic acid extracts,14,15,20 direct spectophotometry is not recommended. A solution is offered by DNA-intercalating compounds such as Hoechst dye 33258, which results in DNA absorbance at different wavelengths.2,11,20 Gel electrophoresis, on the other hand, is often the method of choice as it allows the direct estimation of DNA quantity and quality (degree of shearing, presence of humics). Extracted DNA can be used directly for analysis, or it may be separated into fractions, which are based on, for instance, molecular size or on relative G+C content. To enhance successful detection of a released GMM, van Elsas et al56 isolated the 2-23 kb fraction of their soil DNA, as the specific targets were expected to be prevalent in this fraction. Nusslein and Tiedje63 succeeded in separating soil DNA into high-G+C (63%) and low-G+C (35%) fractions, after which clones generated via PCR with these fractions were sequenced. The former fraction was dominated by Pseudomonas, Rhizobium-Agrobacterium and Rhodospirillum types, whereas the latter had a dominance of Clostridium sequences. Detection of specific sequences in purified DNA extracts can proceed by a number of different strategies, i.e., 1. direct hybridization with a specific probe, 2. DNA labeling and use in probing (reverse sample probing), 3. direct restriction, followed by analysis on an agarose gel and/or cloning and 4. PCR amplification followed by molecular analysis. These analyses and their intricacies are summarized hereunder. 1. Direct hybridization. To directly assess the presence and relative abundance of target sequences in the environment studied, the environmental DNA can be hybridized with labeled probes for these targets, in dilution dot blot or even Southern setups.54,64 For the analysis of target abundancy, calibration of the signals, e.g. via the use of target DNA added in known dilutions, is needed. An advantage of direct hybridization is that it is independent from amplication techniques. Moreover, it allows the direct assessment of rearrangements in the genes of released organisms.65 However, a major disadvantage is its low sensitivity. For an average soil with 108-109 cells per g, it is often difficult to detect targets present at levels below 105 per g. 2. DNA labeling and use in probing. Applications of environmental DNA, in particular DNA from oil fields, include its use as a probe to assess the prevalence of specific targets, in a reverse hybridization setup coined reverse sample probing.66 For this purpose, the DNA should be of sufficient purity to allow (enzymatic) labeling and subsequent hybridization. 3. Direct restriction and cloning. Commonly, restriction enzyme digestion and cloning are performed on previously amplified DNA. However there might be instances in which it is important to obtain a bank of environmental fragments, in particular when the potential biases introduced by PCR are to be bypassed. For that purpose, the DNA should be amenable to cutting with common enzymes, to allow cloning. 4. PCR amplification. The vast majority of analyses performed with environmental DNA are based on the use of PCR amplification, since PCR offers the advantage that the targets, even when present as minority sequences, can be quickly amplified


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to easily detectable or clonable quantities. The potential of this approach is tremendous, and it has revolutionized molecular microbial ecology. In accordance with primer choice, specific target organisms or genes can be detected and quantified,50-51 or broader groups can be fingerprinted by techniques such as denaturing or temperature gradient gel electrophoresis (D/TGGE),67,68 terminal restriction fragment length polymorphism (T-RFLP),69 single strand conformational polymorphism (SSCP)70 or amplified ribosomal DNA restriction analysis (ARDRA). This field is currently exploding, with new applications being reported almost monthly. However, due to problems inherent to mixed community PCR,71 the real value of the resulting fingerprints for actual in situ microbial diversity remains to be assessed. Furthermore, an important step forward in the analysis of D/TGGE profiles is the use of probes based on the variable region 6 (V6 probes) of the 16S rDNA.72 This highly variable region allows for the generation of highly specific probes which can serve to identify bacterial types in gels, or to link the bands from D/TGGE gels to either isolates or cloned 16S community amplicons.

3.6. Extraction and Purification of RNA In contrast to DNA, which serves as a pool of genetic information, RNA has multiple cellular functions. Generally, three classes of RNA can be distinguished, i.e., messenger (m)RNA, ribosomal (r)RNA and transfer (t)RNA. mRNA molecules are transcribed from genes during gene expression. These transcripts are translated into amino acid sequences at ribosomes, which are made up of rRNA molecules and proteins. tRNA molecules serve as carriers for amino acids in this translation process in protein biosynthesis at the ribosomal sites. In microbial ecology, all three classes of RNA have been targeted to characterize structures or functions of microbial communities. Additionally, RNA can be present in the environment in the form of viral genomes. Protocols for the extraction and detection of viral RNA from environmental samples are available73-76 and will not be discussed further in this context. RNA extraction and purification from environmental matrices follow the same lines described for DNA extractions. However, due to the high abundance and persistence of RNases in the environment, RNA extractions generally require more precautions than those of DNA. The use of RNase-inhibiting compounds, like diethylpyrocarbonate (DEPC), chaotropic RNA stabilizing reagent or SDS to inactivate proteins can help to increase extraction efficiencies from environmental samples.78,79 Protocols have been developed to simultaneously obtain different types of RNA or even both RNAs and DNA from environmental samples,6,44,45,80 whereas other protocols aimed at extracting specific RNAs.78 Similar to DNA (see above; Table 3.1), recently developed RNA extraction protocols take advantage of commercially available nucleic acid extraction and purification kits, thereby replacing time- and material-consuming purification steps like ultracentrifugation or dialysis. Limitations of existing protocols are correlated with RNA instability (especially prokaryotic mRNA), and with the co-extraction of contaminating substances, mainly DNA or humic acids. In this regard, soil or sediment samples are more laborious for purification and less sensitive for detection compared to samples of aquatic origin. Therefore, it is not surprising that, in early work, the detection of bacterial transcripts was shown to be several orders of magnitude more sensitive in marine environments81 than in soil.59, 82 Various strategies can be used to obtain suitable material for rRNA analysis, i.e. direct or indirect (cell extraction first) extraction of RNA from environmental samples, or isolation of ribosomes. These approaches have been successfully applied for 16S rRNA based studies. For instance, using analyses based on direct RNA extraction, Hahn et al83 were able to detect poorly culturable Frankia sp. in soil. Felske et al84 characterized the bacterial

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community of a grassland soil on the basis of rRNA obtained from soil-extracted ribosomes. Duarte et al44 generated profiles of soil bacterial communities using reverse transcription and PCR amplification of RNA extracted from the bacterial fraction. The RNA obtained from pre-extracted cells or ribosomes is likely to be less contaminated with coextracted material like humic acids.44,85 However, recent progress in optimizing PCR efficiencies even in the presence of a substantial amount of contaminating humic acids by a considerate selection of heat-stable DNA polymerases and additives like single strand stabilizing proteins, suggests that partially purified RNA might be good targets for RT-PCR analysis.14,55,86 Generally the yield of ribosomal RNA has been reported to be higher with direct or indirect extractions than with the ribosome isolation method.44,87 Early protocols to directly extract total RNA from soil could not detect intact ribosomal RNA molecules,59,88,89 but recently a rapid and direct extraction of intact 5S, 16S, 18S, 23S and 28S RNA, which were amplifiable by RT-PCR has been reported.79

3.7. Detection of RNA 3.7.1. Analysis of Gene Expression on the Basis of mRNA Assessments Analysis of mRNA directly extracted from environmental samples is conducted to investigate habitat dependent regulations of gene expression. In eukaryotic organisms, transcribed mRNA molecules are altered by various co- and posttranscriptional modifications such as capping of the 5' end and polyadenylation of the 3'-end.90 These modifications increase the stability of the molecules to half-life values of several hours. The poly-A 3'-end is a suitable target for the isolation of eukaryotic transcripts by poly-T hybridization using commercially available matrices. In particular, poly-T-coated magnetic beads have been found useful in this respect. Tebbe et al91 extracted mRNA from yeast cells (Saccharomyces cerevisiae, Pichia angusta) in soil by a direct approach, hybridizing poly-T magnetic beads in soil slurries, previously treated for total cell lysis. Transcripts from 10 cells per g soil were detectable after reverse transcription of mRNA and subsequent amplification by PCR. The expression of a recombinant gene was found to become extinct before elimination of the organisms from the environment. A similar polyA-polyT hybridization protocol was developed for the analysis of specific gene expression (e.g. lignin peroxidases) of the white-rot fungus Phanerochaete crysosporium in soil.92-94 In contrast to eukaryotic mRNA, bacterial mRNAs have half-life values in the range of only 0.5-30 min.95 Also, they lack a common target region like the poly-A 3’-end. In order to detect transcripts of biodegradative genes (nahAB transcripts) and the mercury reductase gene merA by gene probe hybridization, over 107 target cells had to be present per g of soil.59,82 Enhanced sensitivity (approximately 2 orders of magnitude) in the detection of bacterial gene expression was achieved by Fleming et al96 who protected specific mRNAs of the napthalene degradation encoding nahA gene by hybridization with antisense RNA in a ribonuclease protection assay. Target cells can also be extracted from soil and gene expression can be analyzed by whole-cell hybridization with digoxigenin-labeled probes. This approach has been used to detect the expression of a protease gene by Bacillus megaterium in soil. The detection limits, however, were above 106 cells g-1 soil.97 Moreover, as indicated before, cells upon extraction from soil may change their physiological status and hence their pattern of gene expression does not necessarily reflect that in situ. Thus, the detection limits for analysis of bacterial gene expression in soil are still several orders of magnitude above the sensitivity desired for most ecological studies. To date, the use of recombinant reporter genes (Chapters 5-7) represents a more sensitive approach to study bacterial gene expression in soil.98,99


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In aquatic samples, the problem of low sensitivity can be alleviated by concentrating bacterial cells prior to RNA extraction.100 In a pioneering study,81 106 cells (concentrated from 100 ml seawater) were necessary to detect the expression of the plasmid-encoded neomycin phosphotransferase gene (nptII) by Vibrio sp. Detection was by hybridization of target mRNA with single-stranded RNA probes. Similar detection limits were observed in studies on the expression of merA by Pseudomonas aeruginosa and microbial communities in lake water.101,102 Quantification of mRNA via direct hybridization or quantitative PCR will be discussed further under section 3.8, below. Differential display (DD) is a PCR-based method which allows to generate characteristic patterns of expressed genes in an organism without a priori sequence information. Recently, this technique has been applied to analyze prokaryotic mRNA following extraction from soil. The expression of a gene (todC1) involved in the degradation of toluene by an inoculated Pseudomonas putida in soil could be demonstrated.103 Additionally, gene expression of the soil microbial community, triggered by the addition of toluene was detected. The DD-technique has the potential to characterize responses of natural microbial communities to, for instance, stress and can eventually help to identify new genes and enzymes of environmental significance.

3.7.2. rRNA-Characterization of Growing Cells in Microbial Communities Analysis of rRNA has become a major technique in the characterization of microbial activities and community structures in the environment.104-106 The various rRNA types contain regions that are conserved among all living organisms as well as regions characteristic for their phylogenetic individuality, offering both targets for PCR and markers for differentiation (Chapter 4). Analysis of rRNA is often preferred over that of DNA, since the number of ribosomes and, thus, rRNA, correlates with the growth rate for a number of bacteria. 107109 Hence, for these types, rRNA analysis would mainly include active organisms and not non-viable or dormant cells. However, other bacterial groups do not seem to shift their ribosome numbers up and down in accordance with their growth rate. Furthermore, the presence of multiple rRNA targets per cell enhances the sensitivity of detection compared to DNA-based detection. Finally, compared to whole genomes, the diversity of sequences is dramatically reduced, and less diverse sequences might be less prone to non-specific primer binding. However, this point is open for debate.110 Using domain-specific gene probes, Ogram et al89 found that rRNA directly extracted from sediment was mainly of bacterial, archaeal and eukaryotic origin. To obtain sufficient numbers of target sequences for the analysis of variable sequences, regions were amplified by PCR using primers which bind to highly conserved sites.111 Such PCR-mediated analysis of rRNA is possible at the DNA level or directly at the RNA level. If RNA is the template, reverse transcription is necessary prior to PCR. Most environmental studies on microbial community structures have focussed on 16S or 18S rRNA, but other regions may become more important during the next years, e.g. the bacterial 23S rRNA112 and the internally transcribed spacer (ITS) regions.113,114 The comparison of 16S or 18S rRNA PCR products generated with community RNA and DNA as templates might provide insight in the ratio of active (growing) and inactive cells in particular environments such as soil.44 Using denaturing or temperature gradient gel electrophoresis (D/TGGE) or single strand conformation polymorphism (SSCP), the intensity of such products can be visualized and compared.44,67,70 Significant differences can then be characterized by DNA sequencing. RNA and DNA based characterizations of fecal communities were similar,115 suggesting that due to the high nutrient content of fecal material, the dominating microorganisms in this system were probably actively growing. In environments with large numbers of inactive cells like bulk soil, however, differences between

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rRNA and DNA based techniques can be more significant, as shown by Duarte et al.44 DGGE analysis of PCR or RT-PCR products generated with either DNA or RNA from the same bacterial fraction showed that, next to similarities in the community profiles, there were clear dissimilarities, suggesting that there were differences in the activity levels between the various bacterial groups underlying the profiles.

3.8. Quantification of Specific Targets in Environmental Nucleic Acids Specific targets in environmental DNA or RNA can be directly quantified by hybridization in a dot blot setup, using careful calibration with a dilution range of the target in the same background.64 Among numerous other applications, the fate in soil of a genetically marked self-transmissible plasmid, RP4, has thus been successfully monitored.56 Quantification of specific mRNA can also be achieved by hybridization with a single-stranded RNA probe. To assess the level of gene expression, the amount of mRNA should be normalized to the gene dose, i.e., the number of genes present in the DNA extracted from an environmental sample. Such an approach has been successfully used by Pichard and Paul116 to study the temperature-dependent expression of a biodegradation gene (xylE) of Vibrio sp. in seawater. However, due to the relative insensitivity of common hybridization protocols,54 probing methods are generally restricted to the assessment of relatively high target numbers present in the sample. Hybridization-based quantification has, thus, been superseeded by PCR- or RT-PCR-based quantitative methods. Quantification via PCR, eventually preceded by a reverse transcription step of mRNA, can be achieved if a relationship between the number of target molecules obtained in the amplification product and the original target number can be established. Several different procedures have been cogitated and applied,117 of which the most important ones are (1) Most-probable-number PCR (MPN-PCR), and (2) Competitive PCR using an internal standard. Other approaches, such as direct quantification,118 quantification by comparison to an external standard or by comparison to a co-amplicon, analysis of PCR kinetics and molecular beacon PCR (in which a fluorescent molecule is formed per successful amplicon produced in the PCR) have also been proposed. These later applications have either met with problems when applied to environmental samples or have found little application yet, and will therefore not be further discussed.

3.8.1. MPN-PCR The principle of MPN-PCR is similar to that in other MPN enumeration procedures. Briefly, the sample material (extract) is serially diluted and amplified, in a number of replicates, to a level where detection is not possible anymore (outdilution). Theoretically, this would be the case at the point of complete removal of all target sequences from the PCR reaction mix. However, in practice the cutoff points can be slightly higher due to inefficiencies in PCR amplification of targets in environmental DNA. The data of all amplification reactions are recorded, and the cutoff levels are determined in the replicates. A table based on statistical principles will then indicate the most probable number of virtual amplifiable targets (VAT) in the dilution assessed, and, thus, in the sample. If the minimal number of targets amplifiable under the conditions used equals 1, then the VAT equals the actual amplifiable numbers; if there is a discrepancy, a compensation factor should be included in the calculations. Nesme et al 119 successfully used MPN-PCR to quantify the number of pathogenic Agrobacterium sp. strains in soils. MPN-PCR has further been employed, for instance, to assess the numbers of targets of Mycobacterium chlorophenolicum, a pentachlorophenoldegrading bacterium, in soil.61 The method was specific and provided positive detection of


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the target down to about 103 targets per g of soil. In addition, the fate of introduced Paenibacillus azotofixans vegetative cells and spores in soil and wheat rhizosphere could be successfully monitored in microcosms using specific MPN-PCR.51 However, an important drawback of MPN-PCR is the excessive number of replicate PCR reactions needed, as well as the careful controls that are required for PCR efficiency. This makes the method rather laborious and expensive. MPN-PCR has now been superseeded by competive PCR methods as discussed below.

3.8.2. Competitive PCR Competitive PCR (cPCR) is based on the principle of co-amplification with the same primers, at presumably similar rates, of known amounts of an easily recognizable added indicator (competitor) molecule with the actual target nucleic acid in the sample. To obtain comparable amplification rates between target and competitor, both the sizes and sequences of the two products should not be widely divergent. Moreover, control of PCR conditions (rate and extent of amplification) is required to avoid biases due to effects of primer:product annealing competition.120 Jansson and Leser117 reviewed the intricacies of competitive PCR, and the reader is referred to this paper for detailed information. In a common competitive PCR assay, a set of reactions is set up with varying amounts of competitor DNA added to the environmental extract containing an unknown amount of the target. Following competitive PCR, the number of targets in the sample can be inferred from the PCR reaction in which the number of indicator molecules equals the number of specific targets. Alternatively, a standard curve is prepared with a dilution series containing a known number of target DNA molecules and a fixed number of competitor DNA molecules. The ratio of the competitor:target band intensities after PCR amplification is plotted versus the original target number. This simplifies the cPCR approach, since a single sample containing an unknown target DNA concentration can be co-amplified together with the competitor DNA at the fixed concentration used for preparation of the standard curve. The target DNA concentration in the sample can be determined from the ratio competitor:target products by interpolation from the standard curve.117,121 There are certain restrictions to the type and size of indicator molecule, as it is absolutely required that amplification rates are similar between competitor and target. The competitor should, for instance, be similar to the target with respects to size, base sequence and flanking sequences. Suitable competitors are artificially constructed or natural DNA fragments of similar size and with similar primer annealing sites but different internal sequence, which are separable on denaturing gradients. For 16S rDNA sequences, competitors can be easily generated via low-stringency PCR with heterologous templates, yielding molecules with similar size yet different sequence to the target. Also, small deletions or insertions can be introduced into the target region, in order to obtain an indicator which is separated by size on an agarose gel.117 Competitive PCR has been successfully applied in studies on the fate of specific bacteria released into the environment. Using amplification of a specific (B13) fragment, Leser et al 122 successfully monitored the decline, as well as the response to a selective agent (4-chlorobenzoate), of the genetically modified Pseudomonas sp. B13 (FR1) in marine microcosms. Also, a competitive quantification system was developed to follow the fate of the pentachlorophenol degrader Sphingomonas chlorophenolica in soil.123 A competitor was fortuitously found when studying sizes of amplicons generated with bacterial primers on different bacterial species, as a product generated with Paenibacillus polymyxa was slightly smaller than the product generated with S. chlorophenolica, while amplification rates were similar. This allowed separation of the two products by electrophoresis through common agarose gels. S. chlorophenolica was thus shown to persist for a long time in pentachlorophenol containing

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soil. cPCR was further found to be a very accurate method for quantitation of a genetically modified cyanobacterium added to Baltic sea sediment.121 In these experiments, the competitor DNA was added to the sediment samples before DNA extraction and PCR amplification in order to account for variations in extraction efficiencies between samples. However, as mentioned, competitive PCR based on amplification of DNA targets does not assess viable cell numbers. Hence, if a focus on active cells rather than total cells is required, the method should be combined with other methodology, viz microscopic assessments of viable cells. The activity (expression) of specific genes can be quantified by using PCR preceded by a reverse transcription step (RT-PCR). This method is very sensitive for the detection of transcripts, as shown with eukaryotic microorganisms;91,92 detection of bacterial gene expression by RT-PCR in activated sludge was possible at a detection threshold of 106 CFU g-1 soil.124 Competitive RT-PCR allows for quantitative determinations of gene expression, as was shown in studies on the expression of peroxidases by Phanerochaete chrysosporium in soil.93,94 Obviously, quantification on the basis of two enzymatic reactions needs a series of controls to compensate for the putative biases of both processes.

3.9. Conclusions and Perspectives for Further Development From a practical perspective, several of the DNA and RNA extraction and analysis methods discussed herein are suitable for the screening of large sample numbers. They are thus useful for the assessment of the fate of GMMs and/or their genes (activity) following introduction into environmental matrices. If required, they also facilitate the detection of naturally occurring DNA or RNA sequences in soil microbes. Due to the current lack of comprehensive data with respect to the suitability of different protocols for different environmental matrices, unfortunately there is no standard nucleic acid protocol that can be recommended for the wide array of environments in which the technology is applicable, not even for soils. However, two recent papers indicated some progress towards a unification of protocols in that amplifiable DNA can be obtained from a range of different soils by the flexible and educated use of different purification routines61,62 On the other hand, real routine and high-throughput use can only be achieved if methods can be kept simple and can be miniaturized or even robotized, as suggested by Kuske et al.20 The great strength of the nucleic acid based detection strategies lies in the fact that direct and cultivation-independent data can be obtained with respect to the numbers (DNAbased strategies) and activities (RNA-based strategies) of microorganisms in complex matrices. The DNA based assessments will often also detect extracellular DNA (the naked genes) as well as DNA from either non-viable, non-culturable, injured or even dead cells. Thus, they can provide a very complete picture of the occurrence of the molecular target under study, which is important when the fate of recombinant genes in the environment is to be monitored. However, it is less relevant to obtain such information when only the viable cells present in the environment are to be targeted. For these studies, DNA-based methods are less suitable, as even with methods that exclusively target microbial cells (carefully controlled for the co-extraction of extracellular DNA), there is no inherent link to viability. Therefore, these methods need a link to molecular or traditional viability assessments, an obvious choice being analyses based on RNA. As a corollary, methods based on RNA suffer less from biases related to the presence of naked DNA, as, due to their presumed rapid decline in most environments, all RNA types tend to reflect cellular activity rather than cellular abundance. Furthermore, even though for instance the bead beater is known to efficiently lyse a major proportion of bacterial species including many gram-positives,50,51 there is no absolute certainty that cell lysis by this method will yield nucleic acid mixtures that are truly representative for the soil bacterial community. For instance, the findings of Moré et al 47


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suggest that lysis may well be confined to the larger cell-size fraction of the microbial community, since the numerous minute or dwarf cells present essentially escaped lysis. On the other hand, the nature and significance of these dwarfs is currently under heavy debate, as their putative role in natural environments is blurred. These considerations have to be taken into account when DNA or RNA extraction protocols are to be used for microbial community structure or fate studies in soil. In addition, the majority of extraction protocols not only yield bacterial, but also eukaryotic DNA, which may originate from a variety of source organisms present in the system (e.g., fungi, protozoans, nematodes and/or plants). For quantification purposes, a eubacterial probe is necessary to determine the proportion of bacterial DNA in the total extracts obtained. A major criticism on studies that were exclusively based on environmental DNA, including those that identified novel microbial species by sequencing of cloned 16S ribosomal amplicons, has been the absence of significant evidence for the cellular nature of these sequences. To bridge the gap between the evidence for novel types exclusively on the basis of clone sequences and knowledge obtained from sound microbiological work in which microbial functioning is central, a link to traditional, isolation-based microbiology has been advocated.125 However, another strategy with potential to link the direct molecular information with that of microbial cells can be based on the detection of cells following their reaction with a labeled probe, a process called whole cell hybridization. This procedure has been applied successfully in aquatic environments, but it has met with problems of signal weakness in soils.126 Recently, it was shown that whole cells from the environment can also be used as targets for PCR (“in situ PCR”) or reverse transcription-PCR,127-129 and this approach has a potentially enhanced sensitivity. This method has been successfully applied to aquatic systems, whereas its application to soil is still awaiting. Both methods have clear potential as complementary tools to support findings obtained on the basis of the extractive protocols. At this point in time, a major drawback may still be their sensitivity to false positive results, as well as the limitations with respect to minimal cell densities needed for successful observation of cells on microscopy slides. However, the combination of molecular fingerprinting methods with the whole cell hybridization approach to monitor the potential enrichment and isolation of “unculturable”, VBNC or “hitherto—as yet—uncultured” bacteria is a powerful tool which has so far been only sparsely explored. Thus, albeit with a definite right of their own, molecular analyses on the basis of extractive approaches clearly gain enormous scientific strength and meaning when they are tied together with methods based on cultivation approaches. It is foreseeable that a polyphasic strategy, in which the extractive, the whole cell hybridization and the cultivation-based approaches are combined, ultimately offers the best perspective for future studies on the fate and activity of microorganisms in their natural habitats.

Acknowledgments This work was supported by grants from the EU-BIOTECH Programme. We thank L.S. van Overbeek for critically reading the manuscript. J.D. van Elsas and C. C. Tebbe are members of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Akkermans ADL, van Elsas JD, de Bruijn FJ. Molecular Microbial Ecology Manual. Kluwer Academic Publishers, Dordrecht, 1995. 2. Trevors JT, van Elsas JD. Nucleic acids in the Environment; Methods and Applications. Springer Verlag, Heidelberg, 1995.

46


3. Van Elsas JD, Duarte GF, Rosado AS et al. Microbiological and molecular biological methods for monitoring microbial inoculants and their effects in the soil environment. J Microbiol Meth 1998; 32:133-154. 4. Torsvik V. Isolation of bacterial DNA from soil. Soil Biol Biochem 1980; 12:15-21. 5. Holben WE, Jansson JK, Chelm BK et al. DNA probe method for the detection of specific microorganisms in the soil bacterial community. Appl Environ Microbiol 1988; 54:703-711. 6. Ogram A, Sayler GS, Barkay TJ. DNA extraction and purification from sediments. J Microbiol Meths 1987; 7:57-66. 7. Steffan RJ, Goksoyr J, Bej AK et al. Recovery of DNA from soils and sediments. Appl Environ Microbiol 1988; 54:2908-2915. 8. Dijkmans R, Jagers A, Kreps S et al. Rapid method for purification of soil DNA for hybridization and PCR analysis. Microb Rel 1993; 2:29-34. 9. Josephson KL, Gerba CP, Pepper IL. Polymerase chain reaction detection of nonviable bacterial pathogens. Appl Environ Microbiol 1993; 59:3513-3515. 10. Malik M, Kain J, Pettigrew C et al. Purification and molecular analysis of microbial DNA from compost. J Microbiol Meth 1994; 20:183-196. 11. Lovell CR, Piceno Y. Purification of DNA from estuarine sediments. J Microbiol Meth 1994; 20:161-174. 12. Purdy KJ, Embley TM, Takii S et al. Rapid extraction of DNA and rRNA from sediments by a novel hydroxyapatite spin-column method. Appl Environ Microbiol 1996; 62: 3905-3907. 13. Cullen DW, Hirsch PH. Simple and rapid method for direct extraction of microbial DNA from soil for PCR. Soil Biol Biochem 1998; 30:983-993. 14. Tebbe CC, Vahjen W. Interference of humic acids and DNA extracted directly from soil in detection and transformation of recombinant DNA from bacteria and a yeast. Appl Environ Microbiol 1993; 59:2657-2665. 15. Smalla K, Cresswell N, Mendonca-Hagler LC et al. Rapid DNA extraction protocol from soil for polymerase chain reaction-mediated amplification. J Appl Bacteriol 1993; 74:78-85. 16. Tsai, Y-L, Olson BH. Rapid method for separation of bacterial DNA from humic substances in sediments for polymerase chain reaction. Appl Environ Microbiol 1992; 58:2292-2295. 17. Tsai Y-L, Olson BH. Detection of low numbers of bacterial cells in soils and sediments by polymerase chain reaction. Appl Environ Microbiol 1992; 58:754-757. 18. Porteous LA, Armstrong JL. A simple method to extract DNA directly from soil for use with polymerase chain reaction amplification. Curr Microbiol 1993; 27:115-118. 19. Porteous LA, Armstrong JL, Seidler RJ et al. An effective method to extract DNA from environmental samples for polymerase chain reaction amplification and DNA fingerprint analysis. Curr Microbiol 1994; 29:301-307. 20. Kuske CR, Banton KL, Adorada DL et al. Small-scale DNA sample preparation method for field PCR detection of microbial cells and spores in soil. Appl Environ Microbiol 1998; 64:2463-2472. 21. Torsvik V, Goksoyr J, Daae FL. High diversity in DNA of soil bacteria. Appl Environ Microbiol 1990; 56:782-787. 22. Paul JH, Pichard SL. Extraction of DNA and RNA from aquatic environments. In: Trevors JT, van Elsas JD, eds. Nucleic acids in the environment. Methods and applications. Berlin: Springer Verlag 1995: 153-177. 23. Bej AK, Mahbubani MH, Dicesare JL et al. Polymerase chain reaction-gene probe detection of microorganisms by using filter-concentrated samples. Appl 9-3534. 24. Fuhrman JA, Comeau DE, Hagström A et al. Extraction from natural planctonic microorganisms of DNA suitable for molecular biological studies. Appl Environ Microbiol 1988; 54:1426-1429. 25. Somerville CC, Knight IT, Straub, WL et al. Simple rapid method for direct extraction of nucleic acids from aquatic environments. Appl Environ Microbiol 1989; 55:548-554.


47

26. Pickup RW, Rhodes G, Saunders JR. Extraction of microbial DNA from aquatic sources: freshwater. In: Akkermans ADL, van Elsas JD, de Bruijn FJ, eds. Molecular Microbial Ecology Manual. Dordrecht: Kluwer Academic Publishers, 1995:1.1.2:1-11. 27. Paul J. Extraction of microbial DNA from aquatic sources: Marine environments. In: Akkermans ADL, van Elsas JD, de Bruijn FJ, eds. Molecular Microbial Ecology Manual. Dordrecht: Kluwer Academic Publishers, 1996:1.1.1:1-13. 28. Dixon B. Biofilms: Diversity in action. ASM News 1998; 64:484-485. 29. Hopkins DW, O’Donnell AG. Methods for extracting bacterial cells from soil. In: Wellington EMH, van Elsas JD, eds. Genetic Interactions Among Microorganisms in the Natural Environment.1992: 104-112. London: Pergamon Press. 30. Torsvik V. Cell extraction method. In: Akkermans ADL, van Elsas JD, de Bruijn FJ eds. Molecular Microbial Ecology Manual. Dordrecht: Kluwer Academic Publishers, 1995: 1.3.1:1-15. 31. Priemé A, Sitaula JIB, Klemedtsson AK et al. Extraction of methane-oxidizing bacteria from soil particles. FEMS Microbiol Ecol 1996; 21:59-68. 32. Leung K, Trevors JT, van Elsas JD. Extraction and amplification of DNA from the rhizosphere and rhizoplane of plants. In: Nucleic Acids in the Environment; Methods and Applications. In: Trevors JT, van Elsas JD, eds. Heidelberg: Springer Verlag,. 1995: 69-87. 33. Strickland TC, Sollins P, Schimel DS et al. Aggregation and aggregate stability in forest and range soil. Soil Sci Soc Am J 1988; 52:829-833. 34. Donegan K, Matyac C, Seidler R et al. Evaluation of methods for sampling, recovery, and enumeration of bacteria applied to the phylloplane. Appl Environ Microbiol 1991; 57:51-56. 35. Ramsay AJ. Extraction of bacteria from soil: Efficiency of shaking or ultrasonication as indicated by direct counts and autoradiography. Soil Biol Biochem 1984;16:475-481. 36. Lindahl V, Aa K, Olsen RA. Effects on microbial activity by extraction of indigenous cells from soil slurries. FEMS Microbiol Ecol 1996; 21:221-230. 37. Faegri A, Torsvik VL, Goksoyr J. Bacterial and fungal activities in soil: Separation of bacteria and fungi by a rapid fractionated centrifugation technique. Soil Biol Biochem 1977; 9:105-112. 38. Riis V, Lorbeer H, Babel W. Extraction of microorganisms from soil: Evaluation of the efficiency by counting methods and activity measurements. Soil Biol Biochem 1998; 30:1573-1581. 39. MacDonald RM. Sampling soil microfloras: Dispersion of soil by ion exchange and extraction of specific microorganisms from suspensions by elutriation. Soil Biol Biochem 1986; 18:399-406. 40. Herron PR, Wellington EMH. New method for extraction of streptomycete spores from soil and application to the study of lysogeny in sterile amended and nonsterile soil. Appl Environ Microbiol 1990; 56:1406-1412. 41. Jacobsen CS, Rasmussen OF. Development and application of a new method to extract bacterial DNA from soil based on separation of bacteria from soil with cation-exchange resin. Appl Environ Microbiol 1992; 58:2458-2462. 42. Lindahl V, Bakken LR. Evaluation of methods for extraction of bacteria from soil. FEMS Microbiol Ecol 1995; 16:135-142. 43. Bakken LR, Lindahl V. Recovery of bacterial cells from soil. In: Trevors JT, van Elsas JD eds. Nucleic Acids in the Environment. Berlin Heidelberg: Springer-Verlag 1995: 13-27. 44. Duarte GF, Rosado AS, Keijzer-Wolters AC et al. Extraction of ribosomal RNA and genomic DNA from soil for studying the diversity of the indigenous microbial community. J Microbiol Methods 1998; 32:21-29. 45. Selenska-Pobell S. Direct and simultaneous extraction of DNA and RNA from soil. In: Akkermans ADL, van Elsas JD, de Brujn FJ eds. Molecular Microbial Ecology Manual 1995; 1.5.1:1-17 46. Moore E, Arnscheidt A, Kruger A et al. Simplified protocols for the preparation of genomic DNA from bacterial cultures. In: Akkermans ADL, van Elsas JD, de Brujn FJ eds. Molecular Microbial Ecology Manual 1999 (in press).

48


47. Moré MI, Herrick JB, Silva MC et al. Quantitative cell lysis of indigenous microorganisms and rapid extraction of microbial DNA from sediment. Appl Environ Microbiol 1994; 60:1572-1580. 48. Volossiouk T, Robb EJ, Nazar RN. Direct DNA extraction for PCR-mediated assays of soil organisms. Appl Environ Microbiol 1995; 61:3972-3976. 49. Nazar RN, Robb EJ, Volossiouk T. In: Akkermans ADL, van Elsas JD, de Bruijn FJ eds. Molecular Microbial Ecology Manual. Kluwer Acad Publ Dordrecht 1996; 1.3.6: 1-8. 50. Briglia M, Eggen RIL, de Vos WM et al. Rapid and sensitive method for the detection of Mycobacterium chlorophenolicum PCP-1 in soil based on 16S rRNA gene-targeted PCR. Appl Environ Microbiol 1996; 62:1478-1480. 51. Rosado AS, Seldin L, Wolters AC et al. Quantitative 16S rDNA-targeted polymerase chain reaction and oligonucleotide hybridization for the detection of Paenibacillus azotofixans in soil and the wheat rhizosphere. FEMS Microbiol Ecol 1996; 19:153-164. 52. Van Elsas JD, Duarte GF, Keijzer-Wolters A et al. Analysis of the dynamics of fungal communities in soil via fungal-specific PCR of soil DNA followed by denaturing gradient gel electrophoresis. Molec Ecol 1999 (in press). 53. Picard C, Ponsonnet C, Paget E et al. Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction. Appl Environ Microbiol 1992; 58:2717-2722. 54. Sambrook J, Fritsch EF, Maniatis T. Molecular Cloning. A Laboratory Manual. Second Edition. 1989. New York: Cold Spring Harbor. 55. Al-Soud WA, Radström P. Capacity of nine thermostable DNA polymerases to mediate DNA amplification in the presence of PCR-inhibiting samples. Appl Environ Microbiol 1998; 64:3748-3753. 56. Van Elsas JD, van Overbeek LS, Fouchier R. A specific marker, pat, for studying the fate of introduced bacteria and their DNA in soil using a combination of detection techniques. Plant Soil 1991; 138:49-60. 57. Myrold DD, Martin KJ, Ritchie NJ. In: Akkermans ADL, van Elsas JD, de Bruijn FJ eds. Molecular Microbial Ecology Manual 1995; 1.3.5:1-9. 58. Young CC, Burghoff RL, Keim LG et al. Polyvinyl pyrrolidene. Agarose gel eletrophoresis purification of poylmerase chain reaction-amplified DNA from soil. Appl Environ Microbiol 1998; 59:1972-1974. 59. Tsai YL, Park MJ, Olson BH. Rapid extraction of mRNA from seeded soils. Appl Environ Microbiol 1991; 57:765-768. 60. Moran MA, Torsvik VL, Torsvik T et al. Direct extraction and purification of rRNA for ecological studies. Appl Environ Microbiol 1993; 59:915-918. 61. Van Elsas, JD, Mantynen V, Wolters AC. Soil DNA extraction and assessment of the fate of Mycobacterium chlorophenolicum strain PCP-1 in different soils via 16S ribosomal RNA gene sequence based most-probable-number PCR and immunofluorescence. Biol Fertil Soils 1997; 24:188-195. 62. Zhou J, Bruns MA, Tiedje JM. DNA recovery from soils of diverse composition. Appl Environ Microbiol 1996; 62:316-322. 63. Nusslein K, Tiedje JM. Characterization of the dominant and rare members of a young Hawaiian soil bacterial community with small-subunit ribosomal DNA amplified from DNA fractionated on the basis of its guanine and cytosine composition. Appl Environ Microbiol 1998; 64:1283-1289. 64. Sayler GS, Layton AC. Environmental application of nucleic acid hybridization. Ann Rev Microbiol 1990; 44:625-648. 65. Jansson JK, Holben WE, Tiedje JM. Detection in soil of a deletion in an engineered DNA sequence by using DNA probes. Appl Environ Microbiol 1989; 55:3022-3025. 66. Voordouw G. Reverse sample genome probing of microbial community dynamics. ASM News 1998; 64:627-633. 67. Muyzer G, de Waal EC, Uitterlinden AG. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 1993; 59:695-700


49

68. Heuer H, Smalla K. Application of denaturing gradient gel electrophoresis and temperature gradient gel electrophoresis for studying soil microbial communities. In: van Elsas JD, Trevors JT, Wellington EMH eds. Modern Soil Microbiology 1997; 353-373. New York, NY : Marcel Dekker. 69. Liu WT, Marsh TL, Cheng H, Forney LJ. Characterization of microbial diversity by determining terminal restriction fragment length polymorphisms of genes encoding 16S rRNA. Appl Environ Microbiol 1997; 63:4516-4522. 70. Schwieger F, Tebbe CC. A new approach to utilize PCR-single-strand conformation polymorphism for 16S rRNA-based microbial community analysis. Appl Environ Microbiol 1998; 64:4870-4876. 71. Suzuki M, Rappe MS, Giovannoni SJ. Kinetic bias in estimates of coastal picoplankton community structure obtained by measurements of small-subunit rRNA gene PCR amplicon length heterogeneity. Appl Environ Microbiol 1998; 64:4522-4529. 72. Heuer H, Hartung K, Wieland G et al. Polynucleotide probes that target a hypervariable region of 16S rRNA genes to identify bacterial isolates corresponding to bands of community fingerprints. Appl Environ Microbiol 1999; 65:1045-1049. 73. Tougianidou D, Botzenhart K. Detection of poliovirus in water by direct isolation of the RNA and hybridization with gene probes. Water Sci Technol 1991; 27:219-222 74. Bej AK. PCR amplification of DNA recovered from the aquatic environment. In: Trevors JT, van Elsas JD, eds. Nucleic acids in the environment; Methods and Applications. Heidelberg: Springer Verlag, 1995; 179-218. 75. Tsai YL, Tran B, Palmer CJ. Analysis of viral RNA persistence in seawater by reverse transcriptase-PCR. Appl Environ Microbiol 1995; 61:363-366. 76. Limsawat S, Ohgaki S. Fate of liberated viral RNA in wastewater determined by PCR. Appl Environ Microbiol 1997; 63:2932-2933. 77. Felske A., Akkermans ADL, de Vos WM. Quantification of 16S rRNAs in complex bacterial communities by multiple competitive reverse transcription—PCR in temperature gradient gel electrophoresis fingerprints. Appl Environ Microbiol 1998; 64:4581-4587. 78. Pichard SL, Paul JH. Extraction of microbial RNA from aquatic sources: Marine environments. In: Akkermans ADL, van Elsas JD, de Brujn FJ, eds. Molecular Microbial Ecology Manual 1996; 1.2.1:1-13. 79. Borneman J, Triplett EW. Rapid and direct method for extraction of RNA from soil. Soil Biol Biochem 1997; 29:1621-1624. 80. Selenska S, Klingmüller W. Direct recovery and molecular analysis of DNA and RNA from soil. Microb Releases 1992; 1:41-46. 81. Pichard SL, Paul JH. Detection of gene expression in genetically engineered microorganisms and natural phytoplancton populations in the marine environment by mRNA analysis. Appl Environ Microbiol 1991; 57:1721-1727. 82. Ogunseitan OA, Delgado IL, Tsai YL et al. Effect of 2-hydroxybenzoate on the maintenance of napthhalene degrading pseudomonads in seeded and unseeded soil. Appl Environ Microbiol 1991; 57:2873-2879. 83. Hahn D, Kester R, Starrenburg MJC et al. Extraction of ribosomal RNA from soil for the detection of Frankia with oligonucleotide probes. Arch Microbiol 1990; 154:329-335. 84. Felske A, Wolterink A, Van Lis R et al. Phylogeny of the main bacterial 16S rRNA sequences in Drentse A grassland soils (The Netherlands). Appl Environ Microbiol 1998; 64:871-879. 85. Felske A, Backhaus H, Akkermans ADL. Direct ribosome isolation from soil. In: Akkermans ADL, van Elsas JD, de Brujn FJ eds. Molecular Microbial Ecology Manual 1998; 1.2.4:1-10. 86. Chandler DP, Wagnon CA, Bolton Jr H. Reverse transcriptase (RT) inhibition of PCR at low concentrations of template and its implications for quantitative RT-PCR. Appl Environ Microbiol 1998; 64:669-677. 87. Felske A, Engelen B, Nübel U et al. Direct ribosome isolation from soil to extract bacterial rRNA for community analysis. Appl Environ Microbiol 1996; 62:4162-4167. 88. Moran MA, Torsvik VL, Torsvik T et al. Direct extraction and purification of rRNA for ecological studies. Appl Environ Microbiol 1993; 59:915-918.

50


89. Ogram A, Sun W, Brockman FJ et al. Isolation and characterization of RNA from low biomass deep-surface sediments. Appl Environ Microbiol 1995; 61:763-768. 90. Cattaneo R. Different types of messenger RNA editing. Annu Rev Genet 1991; 25:71-88. 91. Tebbe CC, Wenderoth DF, Vahjen W et al. Direct detection of recombinant gene expression by two genetically engineered yeasts in soil on the transcriptional and translational levels. Appl Environ Microbiol 1995; 61:4296-4303. 92. Lamar RT, Schoenike B, Vanden Wymelenberg A et al. Quantitation of fungal mRNAs in complex substrates by reverse transcription PCR and its application to Phanerchaete chrysosporium-colonized soil. Appl Environ Microbiol 1995; 61:2122-2126. 93. Bogan BW, Schoenike B, Lamar RT et al. Expression of lip genes during growth in soil and oxidation of anthracene by Phanerochaete chrysosporium. Appl Environ Microbiol 1996; 62:3679-3703. 94. Bogan BW, Schoenike B, Lamar RT et al. Manganese peroxidase mRNA and enzyme activity levels during bioremediation of polycyclic aromatic hydrocarbon-contaminated soil with Phanerochaete chrysosporium. Appl Environ Microbiol 1996; 62:2381-2386. 95. Neidhardt FC, Ingraham JL, Schaechter M. Physiology of the bacterial cell: A molecular approach. Sunderland, Mass: Sinauer Associates Inc., 1990. 96. Fleming JT, Sanseverino J, Sayler GS. Quantitative relationship between naphthalene catabolic gene frequency and expression in predicting PAH degradation in soils at town gas manufacturing sites. Environ Sci Technol 1993; 27:1068-1074. 97. Hönerlage W, Hahn D, Zeyer J. Detection of mRNA of nprM in Bacillus megaterium ATCC 14581 grown in soil by whole-cell hybridization. Arch Microbiol 1995; 163:1814-1821. 98. Heitzer A, Webb OF, Thonnard JE et al. Specific and quantitative assessment of naphthalene and salicylate bioavailability by using a bioluminescent catabolic reporter bacterium. Appl Environ Microbiol 1992; 58:1839-1846. 99. Brazil GM, Kenefick L, Callanan M et al. Construction of a rhizosphere pseudomonad with potential to degrade polychlorinated biphenyls and detection of bph gene expression in the rhizosphere. Appl Environ Microbiol 1995; 61:1946-1952. 100. Paul JP, Pichard SL. Extraction of DNA and RNA from aquatic environments. In: Trevors JT, van Elsas JD eds. Nucleic Acids in the Environment; Methods and Applications. Heidelberg: Springer Verlag 1995: 153-177. 101. Jeffrey WH, Nazaret S, Von Haven R. Improved method for recovery of mRNA from aquatic samples and its application to detection of mer expression. Appl Environ Microbiol 1994; 60:1814-1821. 102. Nazaret S, Jeffrey WH, Saouter E et al. merA gene expression in aquatic environments measured by mRNA production and Hg(II) volatilization. Appl Environ Microbiol 1994; 60:4059-4065. 103. Fleming JT, Yao WH, Sayler GS. Optimization of differential display of prokaryotic mRNA: application to pure culture and soil microcosms. Appl Environ Microbiol 1998; 64:3698-3706. 104. Amann RI, Ludwig W, Schleifer KH. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Rev 1995; 59:143-169. 105. Von Wintzigerode F., Göbel UB, Stackebrandt E. Determination of microbial diversity in environmental samples: pitfalls of PCR-based rRNA analysis. FEMS Microb Rev 1997; 21:213-229. 106. Head IM, Saunders JR, Pickup RW. Microbial evolution, diversity, and ecology: a decade of ribosomal RNA analysis of uncultured microorganisms. Microb Ecol 1998; 35:1-21. 107. Wagner R. The regulation of ribosomal RNA synthesis and bacterial cell growth. Arch Microbiol 1994; 161:100-106. 108. Giovannoni SJ, Britschgi TB, Moyer CL et al. Genetic diversity in Sargasso sea bacterioplankton. Nature 1990; 345:60-63. 109. Binder BJ, Liu YC. Growth rate regulation of rRNA content of a marine Synechococcus (Cyanobacteria) strain. Appl Environ Microbiol 1998; 64:3364-3351.


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110. Fuhrman JA, Lee SH, Masuchi Y et al. Characterization of marine prokaryotic communities via DNA and RNA. Microb Ecol 1994; 28:133-145. 111. Devereux R, Willis SG. Amplification of ribosomal RNA sequences. In: Akkermans ADL, van Elsas JD, de Brujn FJ eds. Molecular Microbial Ecology Manual. Dordrecht, NL: Kluwer Acad Publ 1995; 3.3.1:1-11. 112. Ludwig W, Dorn S, Springer N et al. PCR-based preparation of 23S rRNA-targeted groupspecific polynucleotide probes. Appl Environ Microbiol 1994; 60: 3236-3244. 113. Normand P, Ponsonnet C, Nesme X et al. ITS analysis of prokaryotes. In: Akkermans ADL, van Elsas JD, de Brujn FJ eds. Molecular Microbial Ecology Manual Dordrecht, NL: Kluwer Acad Publ 1996; 3.4.5:1-12. 114. White TJ, Bruns T, Lee S et al. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis MA, Gelfand DH, Sninsky JJ, White TJ eds. PCR protocols. A guide to methods and applications. San Diego: Academic Press 1990: 315-322. 115. Zoetendal EG, Akkermans ADL, De Vos WM. Temperature gradient gel electrophoresis analysis of 16S rRNA from human fecal samples reveals stable and host-specific communities of active bacteria. Appl Environ Microbiol 1998; 64:3854-3859. 116. Pichard SL, Paul JH. Gene expression per gene dose, a specific measure of gene expression in aquatic microorganisms. Appl Environ Microbiol 1993; 59:451-457. 117. Jansson J, Leser T. Quantitative PCR of environmental samples. In: van Elsas JD, Trevors JT, Wellington EMH eds. Molecular Microbial Ecology Manual Kluwer Acad. Publ. Dordrecht, NL 1996; 2.7.4:1-19. 118. Romanowski G, Lorenz MG, Wackernagel W. Use of polymerase chain reaction and electroporation of Escherichia coli to monitor the persistence of extracellular plasmid DNA introduced into natural soils. Appl Environ Microbiol 1993; 59:3438-3446. 119. Nesme X, Picard C, Simonet P. Specific DNA sequences for detection of soil bacteria. In: Trevors JT, van Elsas JD eds. Nucleic acids in the environment; Methods and Applications. Heidelberg: Springer Verlag 1995; 111-140. 120. Polz MF, Cavanaugh CM. Bias in template-to-product ratios in multitemplate PCR. Appl Environ Microbiol 1998; 64:3724-3730. 121. Moller A, Jansson JK. Quantification of genetically-tagged cyanobacteria in Baltic sea sediment by competitive PCR. Biotechniques 1997; 22:512-518. 122. Leser TD, Boye M, Hendriksen NB. Survival and activity of Pseudomonas sp. strain B13 (FR1) in a marine microcosm determined by quantitative PCR and an rRNA-targeting probe and its effect on the indigenous bacterioplankton. Appl Environ Microbiol 1995; 61:1201-1207. 123. Van Elsas JD, Rosado AS, Wolters AC et al. Quantitative detection of Sphingomonas chlorophenolica in soil via competitive polymerase chain reaction. J Appl Microbiol 1998; 85:463-471. 124. Selvaratnam S, Schoedel B, McFarland BL et al. Application of reverse transcriptase PCR for monitoring expression of the catabolic dmpN gene in a phenol-degrading sequencing batch reactor. Appl Environ Microbiol 1995; 61:3981-3985. 125. Liesack W, Janssen PH, Rainey FA et al. Microbial diversity in soil: the need for a combined approach using molecular and cultivation techniques. In: van Elsas JD, Trevors JT, Wellington EMH, eds. Modern Soil New York: Microbiology Marcel Dekker, Inc. 1997; 375-439. 126. Hahn D, Amann RI, Ludwig W et al. Detection of micro-organisms in soil after in situ hybridization with rRNA-targeted, fluorescently labeled oligonucleotides. J Gen Microbiol 1992; 138: 879-887. 127. Hodson RE, Dustman WA, Garg RP et al. In situ PCR for visualization of microscale distribution of specific genes and gene products in prokaryotic communities. Appl Environ Microbiol 1995; 61:4074-4082. 128. Chen F, Gonzalez JM, Dustman WA et al. In situ reverse transcription, an approach to characterize genetic diversity and activities of prokaryotes. Appl Environ Microbiol 1997; 63: 4907-4913. 129. Tani K, Kurokawa K, Nasu M. Development of a direct in situ PCR method for detection of specific bacteria in natural environments. Appl. Environ Microbiol 1998; 64:1536-1540.

CHAPTER 4

Detection of Bacteria by Their Intrinsic Markers Éva Tas and Kristina Lindström

4.1. Introduction

T

he detection and identification of specific bacterial species and strains has long been of interest for different applications in agriculture, the food industry, the bioleaching industry and the biological degradation of toxic wastes (bioremediation). In addition, basic and applied sciences, such as microbial ecology, are investing more research into exploring the components of microbial populations living in various ecological niches. Classical microbial identification methods are primarily based on morphology, growth characteristics and metabolic properties of the organism under study. For pathogens or symbiotic organisms, host specificity is also an important feature. In addition, bacteriophage typing, antibiotic resistance, lipopolysaccharide and protein profiles, serological properties, and plasmid profiles of bacteria can be characterized. More advanced immunological techniques include detection of antigens by enzyme-linked immunosorbent assay (ELISA) or by fluorescent antibody labeling. The problem with most of these techniques is that they require microbiological cultivation. Thus, for example, detection of released organisms may not be possible when their numbers are low relative to those of the indigenous population.1 In addition, disadvantages of the immunological methods lay in possible cross-reactions and nonspecific binding. The use of monoclonal antibodies can be a solution,2,3 but the antigens to be detected can be variable depending on growth conditions. Along with the traditional identification methods described above, molecular methods, including detection of specific nucleic acid sequences or profiles are now available. Nucleic acid based detection techniques overcome some of the problems inherent in traditional identification methods (see above) and they are usually more rapid and sensitive than traditional approaches. Detection of specific DNA or RNA sequences in environmental samples enables reliable detection of virtually any group of microorganisms, including those in nonculturable states.4 In particular, DNA:DNA hybridization5 and the polymerase chain reaction (PCR)6,7 are indispensable tools for detection of microorganisms in environmental samples. The reader is also referred to Chapter 3 for details on extraction and analysis of nucleic acids from environmental samples. The targets for molecular identification methods can be intrinsic markers or introduced markers. An intrinsic marker can be defined as a natural (i.e., nonintroduced) DNA sequence or phenotype that serves as a signature for a particular organism or group of organisms. Detection of intrinsic markers is an important tool for identification of specific Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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microorganisms in the environment and complements the use of marker genes for genetic tagging of microorganisms (see Chapters 5-7 describing introduced marker genes). In this chapter we will introduce some examples of the use of intrinsic DNA markers for monitoring of specific microorganisms. We will also describe methods for designing specific nucleic acid probes and PCR primers, which are prerequisites for the application of these detection techniques.

4.2. Brief Description of PCR-Based Fingerprinting Methods PCR based fingerprinting methods are useful for bacterial identification and, as such, are gaining an important role in ecological studies. Some of these methods, such as arbitrarily primed PCR8 and random primed amplified polymorphic DNA (RAPD) analysis,9 are based on amplification of multiple DNA fragments from genomic DNA with random or arbitrary primers, which will result in a characteristic pattern of PCR products for each different organism. Amplified fragment length polymorphism (AFLP) is a variation of these techniques in which genomic DNA is digested with a restriction endonuclease, ligated to adapters, and PCR amplified using primers complementary to the adapters.10 Specific genomic fingerprints can also be achieved by amplification of conserved repetitive bacterial sequences such as Repetitive Extragenic Palindromes (REP), Enterobacterial Repetitive Intergenic Consensus (ERIC) sequences and BOX elements found mainly in gram positive bacteria. 11,12 An example of REP PCR fingerprinting is described in section 4.4.4 below. Other techniques can be used to detect different microorganisms in mixed populations. For example, microbial community diversity can be analyzed by PCR amplification of DNA encoding 16S rRNA genes and separating the products by temperature or denaturing gradient gel electrophoresis (TGGE and DGGE). The resulting fingerprints are indicative of the major species present in a given ecosystem and are sensitive indicators of changes in the structures of microbial communities with different treatments.13

4.3. Development of Specific Hybridization Probes and PCR Primers The first crucial step for both DNA/RNA hybridization and PCR techniques is to find appropriate probe or primer sequences. Fortunately, the development of recombinant DNA techniques and the large amount of sequences accessible in nucleic acid databanks now allow probes and primers to be designed with desired specificity, e.g., for strains, species, genera or other groups. Here we outline a few different approaches for probe design.

4.3.1. Detection of Specific Genes Numerous bacterial genes or gene fragments have been cloned and sequenced, and their occurrence in different organisms is often known. Therefore, these DNA sequences can serve as potential targets for DNA hybridization or PCR. Bacteria that are genetically characterized are often identified using these methods in clinical diagnostics, the food industry and for environmental applications, such as bioremediation.14-21 For example, an intrinsic marker was used for detection of a biodegradative Pseudomonas Burkholderia cepacia strain in soil.22 The intrinsic sequence was a repeated 1.3 kb sequence present in 15-20 copies per cell. This sequence could be detected in sediment samples by dot blot DNA hybridization and PCR, even when the bacterium was no longer detectable by plating methods. In a similar approach, Raaijmakers et al 23 used a siderophore receptor gene as a specific wild type marker for a Pseudomonas putida strain. Engineered marker genes may also be detected by hybridization or PCR (see Chapter 3).19,22,24-29 These techniques are of particular importance for the detection of nonexpressed genetic markers. For example, if a marker gene is repressed or otherwise inactive under certain conditions, nucleic acid probes can still help to detect the gene. Pillai and Pepper30 found that

Detection of Bacteria by Their Intrinsic Markers

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an antibiotic resistance marker carried by Tn5 was not expressed in some cells under high temperature stress. Another example is when the marker is not a fully functional gene but a recombinant fragment detectable by DNA techniques. Van Elsas et al28 used a small eukaryotic DNA fragment, pat from Solanum tuberosum, as a marker to study the fate of Pseudomonas fluorescens introduced into soil. This DNA sequence was not present in indigenous soil bacteria, so problems with background were eliminated.31 Although not expressed as a phenotypic marker, the pat fragment was readily detected by DNA hybridization and by PCR.

4.3.2. Detection of Ribosomal Genes Probes derived from 16S or 23S rRNA genes, or the intergenic transcribed spacer sequence (ITS) separating them, are often used for detection and identification of bacteria. Ribosomal RNA genes are appropriate target sequences against which DNA probes or primers can be directed, and there are several advantages to this approach. Firstly, ribosomal genes are universal and relatively well conserved, but they contain highly conserved and less conserved regions (including ITS), facilitating the design of probes with various specificities.32 Ribosomal RNA is a naturally amplified target (up to 105 copy per cell) that allows in situ detection of single cells by hybridization.33 In addition, ribosomal sequences are well documented in nucleic acid databanks which significantly enhances the potential of probe design. Finally, the probes based on ribosomal sequences are usually oligonucleotides that are easy to synthesize and label. Ribosomal RNA-targeted probes provide important tools for studying the microbial community structure of natural and man-made environments,34-37 but their specificity is limited. Generally, 16S rRNA-derived probes are reliable at the level of genus and above,38 whereas 23S rRNA- and ITS-derived probes distinguish closer relatives such as species and even strains.39 However, phenotypically divergent bacteria can have identical or nearly identical rRNA sequences. In contrast, significant variation in rRNA sequences has been described within single species or between different copies of the rrn operons of a strain.40 These factors should be considered when hybridization probes or PCR primers based on ribosomal sequences are to be used for monitoring.

4.3.3. Use of Randomly Cloned Fragments as Specific Probes When designing hybridization probes, it is not essential to know the function of the selected DNA fragment(s). Randomly cloned DNA fragments, such as entire or enriched genomic libraries, or sub-fragments from partially described genomic regions can be screened for unique sequences.41 Alternatively, specific sequences can be isolated by differential comparison of two or more genomes. Subtraction hybridization, which will be discussed later in more detail, is one method based on this idea. Another potential approach could exploit arbitrarily primed PCR, similarly to differential display which is a method for isolation of genes differentially expressed in different eukaryotic cells or tissues, as described by Liang and Pardee.42 These authors used a set of arbitrary primers for PCR amplification of cDNA generated by reverse transcription from mRNA. They could amplify and clone differentially expressed genes by comparison of the electrophoretic patterns of the amplified cDNA products from different cells. Taking a similar approach, a comparison of the patterns of fragments amplified from genomic DNA from different organisms, by any PCR fingerprinting method mentioned in section 4.2.1, can lead to isolation of DNA fragments unique for a particular bacterial group. The drawback of these methods is that the function, stability and genetic variability of the selected sequences are unknown. On the other hand, these methods can be very useful for rapid preparation of specific hybridization probes, without the requirement for knowledge of sequence information.

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4.3.4. Specificity of Selected Probes and Primers Specificity of each probe and primer pair has to be experimentally evaluated, especially against closely related nontarget groups of bacteria or bacteria inhabiting the environment studied. Since the probes are based on the sequences of strains from culture collections, we cannot be sure that they recognize all possible organisms from an environmental sample that belong to a target group. For example, the developed probes or primers might not detect organisms of environmental importance, which have never been previously cultured. In contrast, unspecificity may be revealed during the environmental application of new probes. This is illustrated by the work of Cullen et al, who used PCR to monitor Rhizobium inoculants in the field.1 One of their markers specifically detected the inoculated strain, while DNA sequences of indigenous bacteria were not amplified. With another set of primers they detected homologous sequences that were present in the indigenous soil microbial population. Some of these nonspecific products comigrated in the gel with the targeted DNA bands, but they could be distinguished by DNA hybridization. Some of the problems with unspecificity could be avoided by increasing the stringency of the reaction conditions used in the assay.

4.3.5. Sensitivity of Detection Detection limits of bacterial strains in environmental samples using the polymerase chain reaction are mainly dependent on the success of nucleic acid extraction. When analyzing aquatic samples, the main problem is the need for filtration of large volumes of water to obtain sufficient cell concentrations. With soil samples, PCR-inhibitory contaminants coextracted with the DNA cause a major problem. Variation in cell lysis and DNA recovery efficiencies could effect not only the yield, but also the ratio of target DNA to nontarget DNA in the extracts used as templates in the PCR reactions. (See chapter 3 for details.) It is possible to increase the sensitivity to a certain extent by fractionating the DNA on CsCl gradients according to its G+C content. This way the target DNA can be enriched in one of the fractions relatively to total environmental DNA.1 Optimization of the PCR reaction conditions, including template concentration, magnesium concentration and number of PCR cycles are ultimate requirements for optimum sensitivity. In some applications special additives in the reaction mixture such as formamide, BSA or betaine can increase both the specificity and the yield of PCR products. As mentioned above, combined use of PCR and hybridization increases not only the specificity but also the sensitivity of detection. For example, in hybridization experiments with mRNA targeted gene specific probes, relatively high cell numbers (> 106 per g of soil or 106 per liter of water) are needed for reliable quantitative detection, although even single cells can be visualized. When PCR is involved in the procedure, 1-100 cells per gram of sediment or soil can be detected.1,22,29,37 However, the sensitivity of detection is always dependent on several factors such as nucleic acid extraction, copy number of the target, assay conditions, probe labeling techniques, and therefore should be considered case specific. Alternatively, nested PCR is another sensitive detection method. This technique uses a second set of primers, which bind to specific sequences internal to the first PCR product. Use of nested PCR confirms the identity of the PCR products and simultaneously increases the sensitivity of detection of the assay.

4.4. Case Study: Specific Detection of Rhizobium galegae Rhizobium bacteria are continuously being released because of their nitrogen fixing capacity that potentially promotes the growth of legumes with which rhizobia live in symbiosis. The Rhizobium-legume symbiosis has been widely investigated and the history of legume fields and inoculations are in many places well documented. Inoculation with new Rhizobium strains is often needed because of the lack of or inefficiency of the indigenous


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rhizobial population. However, inoculation often fails because Rhizobium populations present in the soil may outcompete the newly introduced strain.43 That is why rhizobia are among the most important targets of genetic engineering, aiming to increase nitrogen fixation capacity or inoculation competitiveness.44 As a consequence of genetic engineering and environmental release, sensitive monitoring methods to detect rhizobia in the environment have become essential. Examples of field releases of genetically modified rhizobia are given in Chapters 8-11. Rhizobium galegae is a recently described and well-studied species45 that nodulates Galega officinalis and Galega orientalis (goat’s rue). The latter species is a perennial legume with potential agricultural importance. R. galegae has not yet been genetically engineered, but antibiotic resistant strains have been selected to facilitate laboratory studies. The species has unique taxonomy and host specificity45-47 and it is not indigenous in Finnish soils.48 These properties make it a suitable model organism for studying the fate of inoculant rhizobia in the field in Finland. R. galegae species- and strain-specific probes and PCR primers were developed in our laboratory. They allow specific detection of R. galegae from environmental samples as will be demonstrated in this chapter.

4.4.1. Development of a R. Galegae Species-Specific Probe by Testing Randomly Cloned Fragments As we described above, some approaches for isolation of specific probes comprise screening of partially described regions of a bacterial genome. Since R. galegae is peculiar in that it nodulates only a specific legume host,approaches were adapted to locate R. galegae speciesspecific hybridization probes from the genes determining host specificity. Limited information on the localization of these genes was known from previous hybridization experiments.49 Fragments of DNA, carrying the host specificity gene region of R. galegae, were analyzed for specificity in dot blot hybridization experiments.50 The blots contained total DNA from representatives of different rhizobial species and other soil bacteria. One of the probes hybridized strongly with DNAs from all tested R. galegae strains, but showed a weak reaction with two nontarget organisms (Rhizobium loti and Erwinia carotovora). Therefore, smaller sub-fragments of the probe were further analyzed (Fig. 4.1). One of the sub-fragments showed species-specificity: in Southern blot experiments with EcoRI digested total DNA of different bacteria, as it hybridized only with R. galegae DNA. The DNA fragment was sequenced and PCR primers were designed at the ends of it (Fig. 4.1). The specificity of the primers

Fig. 4.1. Fragments of genomic DNA from R. galegae HAMBI 1174, which carry host specificity genes, were analyzed for species specificity. The 2.4 kb EcoRI-fragment shown here was a good candidate but it weakly hybridized with DNA from nontarget organisms. Its three subfragments were also analyzed, and the 0.9 kb EcoRI–SalI fragment was species specific. Arrows show PCR primers.

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was verified in PCR reactions with purified genomic DNA from an assortment of strains of R. galegae, other rhizobia and some other species.51 PCR amplification always produced an 850-bp specific fragment with R. galegae DNA. PCR-RFLP of this species-specific fragment revealed a difference between R. galegae strains according to their host plant G. orientalis and G. officinalis (Fig. 4.2).

4.4.2. Development of Strain-Specific Probes by Subtraction Hybridization Subtraction hybridization is an efficient method for isolation of DNA present in one genome (probe) but absent in another, otherwise very similar one (subtracter).52-54 Essentially, the procedure involves DNA:DNA or RNA:DNA hybridization carried out either in solution or at a surface. After hybridization, subtracter DNA and probe sequences homologous to it are separated from the mixture, which becomes enriched in nonhybridizing potentially unique target fragments. The hybridization and separation steps can be repeated several times using fresh pools of subtracter DNA in each cycle. The separated DNA pool enriched in specific sequences can be subsequently cloned for further screening or used directly as a probe. DNAs of different strains can be pooled for use as subtracter DNA. Thus, the desired probe specificity can be achieved by controlling the composition of the subtracter DNA.55 A schematic representation of the method is shown in Figure 4.3. The most important factors that affect the success of subtraction or number of cycles needed for effective enrichment are the level of homology the compared genomes share, the stringency of hybridization, and the extent of modifications of the hybridization partners such as restriction enzyme digestion, sonication, and biotin labeling. The subtraction method has several applications such as isolation of tissue-specific or developmental stage-specific genes,52,56 or cloning a gene missing in a deletion mutant.53,57 Subtraction hybridization was also successfully applied for isolation of specific bacterial probes. Welcher et al used the method for the first time and selected specific genomic sequences from Neisseria gonorrhoeae.52 Schmidhuber et al found Streptococcus oralis-specific probes with the same method.41 Bjourson and Cooper successfully isolated strain-specific probes for three different Rhizobium loti strains,54 and Bjourson et al 58 developed an improved method for isolation of strain-specific probes for Rhizobium leguminosarum bv. trifolii.58 In latter work the same researchers developed group-specific DNA probes for R. leguminosarum bv. phaseoli and R. tropici.55 In our laboratory, a R. galegae strain-specific probe was isolated59 by the subtraction protocol of Straus and Ausubel.53 Darrasse et al have applied the same subtraction strategy independently for isolation of probes specific for Erwinia carotovora subsp. atroseptica.60 Figure 4.3 illustrates the subtraction strategy used in our laboratory. The specificity of the selected probe was tested, similarly to the species-specific probe, in numerous dot blot and Southern blot experiments. The fragment was then sequenced, and PCR primers amplifying an internal fragment were designed. The probe and the primers recognized only the strain R. galegae HAMBI 1174, which had been used in the subtraction procedure.

4.4.3. Application of the R. galegae Specific Probes and Primers for Monitoring To study the applicability of detection with strain- and species-specific sequences, a greenhouse experiment was conducted.51 Soil devoid of R. galegae was inoculated with different levels of R. galegae HAMBI 1207, which forms an ineffective symbiosis with G. orientalis. Then seeds of G. orientalis, inoculated with the effective strain R. galegae HAMBI 1174, were sown into the soil. The competition between the two Rhizobium strains was assessed by measuring plant dry weights from the plots and inspecting nodules formed on the plant roots. Multiple PCR with the strain- and species-specific primers were used


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Fig. 4.2. PCR-RFLP of the R. galegae-specific fragment from genomic DNA of different R. galegae strains with the restriction enzymes HinfI and MspI. The host plant is shown above the samples. Lanes: M, molecular weight marker; 1, reference strain, R. galegae HAMBI 1174; 2-15, other R. galegae strains.

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Fig. 4.3. Isolation of a strain-specific probe by subtraction hybridization. HAMBI 1174 and HAMBI 1207 are genetically closely related strains of R. galegae.

directly from crushed nodules (Fig. 4.4). The results were in good agreement with those of dilution platings on selective media, in which the two strains could be counted. The specific primers were also used for the amplification of DNA extracted from soil and peat.61 After developing a suitable method for DNA extraction and purification from the peat samples (for methods see Chapter 3), the PCR assay proved to be very useful for quality control (i.e., confirmation of the inoculum strain) of commercial peat inoculants.


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Fig. 4.4. PCR amplification of the R. galegae species-specific fragment (upper band) and the R. galegae HAMBI 1174 strain-specific fragment (lower band) from crushed nodules. Lanes: λ, molecular weight marker; 1, R. galegae HAMBI 1174 genomic DNA; 2, R. galegae HAMBI 1207 genomic DNA; 3-26, nodules from G. orientalis inoculated with R. galegae; -, negative control without template DNA. Reprinted with permission from Tas É, Leinonen P, Saano A et al. Appl Environ Microbiol 1996; 62:529-535.

4.4.4. Use of REP-PCR for Fingerprinting of Rhizobium galegae DNA primers corresponding to repetitive extragenic palindromic (REP) and enterobacterial repetitive intergenic consensus (ERIC) sequences can be used to produce specific fingerprints of different bacterial genomes.12 These primers were used to fingerprint the genomes of several R. galegae strains.62 For each strain tested, highly reproducible fingerprint patterns were obtained from pure bacterial genomic DNA. The patterns were specific for each strain and REP-PCR grouped the strains according to their host plant, G. orientalis and G. officinalis. The methods were useful for identification of bacteria from pure cultures as well as from Galega root nodules (Fig. 4.5). However, these methods are not suitable for the analysis of mixed bacterial populations such as those, for example, found in soil or peat samples, as the patterns obtained would become too complex.

4.5. Future Prospects Nucleic acid based methods for assessment of the environmental fate and effects of bacteria are under continuous development. In the future, growth and wider availability of nucleic acid sequence databases, further improvement and optimization of molecular techniques as well as application of microarray chips will facilitate monitoring studies. DNA databases are of great help in probe and primer design. Since data are accumulating very fast in nucleic acid sequence databanks, special databases of probes and PCR primers are of great value. There are probe databases accessible via the Internet. Alm et al have established an oligonucleotide probe database.63 Their probes and primers, mainly based on rRNA sequences, are widely used in microbial ecology and environmental microbiology. The authors also provide advice on probe and PCR primer design and characterization. A more complete database for ribosomal sequences and for data analysis tools is maintained at the Center for Microbial Ecology at Michigan State University.64 Another database is available at the Technical University of Munich (http://www.mikro.biologie.tu-muenchen.de). One of the most promising techniques is the direct detection and quantification of gene transcripts (mRNA) by reverse transcription-PCR, which allows assessment of micro-

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Fig. 4.5. REP PCR fingerprint patterns of R. galegae DNA obtained by sonicating root nodules. Lanes: S, lambda BstEII; 1, genomic DNA from reference strain R. galegae HAMBI 540; 2, field nodule (R. galegae HAMBI 540); 3-8, nodules from G. orientalis inoculated with different R. galegae strains; 9, negative control without template DNA. Reprinted with permission from Nick G, Lindström K. System Appl Microbiol 1994; 17:265-273.

bial activities in the field (For a review see ref 65). The RT-PCR technique consists of synthesis of DNA from RNA by reverse transcription, and amplification of a specific DNA fragment by PCR. The method is very sensitive, and by using appropriate controls the detection can be made quantitative. The first environmental applications included detection of food-borne pathogens,66 and microorganisms used in bioremediation of contaminated soils.65 For the selection of a suitable target gene for RT-PCR several characteristics should be considered: the target gene should be abundantly expressed throughout the growth cycle of the organism tested, and its expression should not be regulated at the transcriptional level. Since bacterial mRNAs have a very short half-life, these methods discriminate living (i.e., metabolically active) cells from dead or dormant ones, although DNA from the latter ones can still be detected by DNA-targeted Southern hybridization or amplified by conventional PCR. (Detection of active contra dormant cells is discussed in more detail in Chapter 1.) Some technical difficulties arise from the need for rapid isolation of undegraded mRNA. However, in situ techniques are now under development for the amplification of gene transcripts inside microbial cells.18 In addition, the fingerprints achieved by the PCR-based applications mentioned in section 4.2.1 can be used for identification of microorganisms in pure cultures or relatively simple microbial communities. In highly diverse microbial systems such as soil, the banding patterns are too complex to be assigned to specific organisms but a comparison of microbial communities (diversity as well as quantitative aspects)67 is possible. Particularly the TGGE technique seems to be appropriate for monitoring effects of environmental disturbances such as pollution or release of new organisms.68 Hybridization of RNA and DNA on microarrayed ‘chips’ (on glass or silica surfaces) is another technique that will probably be broadly applied in the near future. Cheng et al prepared nucleic acids of E. coli and other bacteria from human blood and hybridized them with appropriate probes on microchips.69 Since the hybridization of RNA on microarrayed


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chips allows monitoring of gene expression, this technique may prove to be useful in studying the dynamics of cell populations in environmental samples. Thus, microarrays have great potential in medical diagnostics, food monitoring, water testing, and other fields. New approaches based on specific molecular recognition mechanisms are also under development, such as nucleic acid probes detecting proteins and other molecules,70 or synthetic DNA mimics that behave like DNA oligonucleotides but bind to nucleic acids with higher specificity and affinity.71 Drolet et al have developed an ELISA-like assay using an oligonucleotide probe which binds to a protein target.70 A method called SELEX (systematic evolution of ligands by exponential enrichment)72 allowed rapid selection of oligonucleotides that preferentially bind to the target molecule from a population of random sequences. High affinity oligonucleotide ligands could be developed in this way for virtually any target molecule and used, for example, as therapeutic or diagnostic agents.73 Nucleic acid ligands would have several advantages over antibodies.70 For example, they can be easily synthesized and thus accurately replicated with the same binding properties, while antibodies may differ when generated in different laboratories and are also subject to animal-to-animal variation. Secondly, animals are not required for the synthesis of oligonucleotide ligands. Also, the ligands can be generated even for toxic or nonimmunogenic targets. Their small size can also be an advantage for some applications. In the future, further DNA ligands will be developed for diagnostic purposes and may be used for more general approaches. If successful, the method could challenge, for example, in situ hybridization and immune detection methods. An interesting new approach is based on the use of a synthetic molecule peptide nucleic acid (PNA). This molecule has recently been proposed to expand the applications of oligonucleotides.71 PNAs are DNA analogs with a polyamide backbone substituted with purine and pyrimidine base sidechains. The specificity of PNA in binding to complementary nucleic acids is higher than that of DNA strands, and the resulting PNA/DNA and PNA/RNA duplexes have high stability compared to DNA/DNA and DNA/RNA duplexes. Similarly to DNA, PNA can be labeled by biotin, fluorescein or reporter enzymes. Therefore, PNA is a good candidate for hybridization studies and may, one day, share the burgeoning field of DNA-based diagnostics with traditional nucleic acid probes. Along with the above described sophisticated genetic approaches, phenotypic description of microbial communities is equally important. Metabolic fingerprints of individual organisms or microbial communities can be determined by the BIOLOG‘ system, which colorimetrically detects the utilization of various carbon substrates. Phospholipid fatty acid (PLFA) profiles are also increasingly being used to characterize the structure of microbial communities.74,75

4.6. Conclusion To assess the survival and behavior of deliberately released strains in the environment, specific markers are required for monitoring the populations of both the released and the indigenous strains. Intrinsic markers are of great value in these studies and their detection complements the use of tagging genetically introduced marker genes. In addition, some applications, e.g. detection of pathogens or food-borne microorganisms, do not allow the use of genetically engineered markers. Therefore, detection techniques based on specific intrinsic markers are indispensable. This chapter focused on these techniques and discussed different methods for selection of suitable molecular probes and primers. In addition to exploring previously described genes or ribosomal sequences, an alternative for probe selection is screening randomly cloned fragments. This approach can provide specific probes also for organisms for which sequence data are limited. It was illustrated in this chapter by the isolation of strain- and species-specific probes from R. galegae. The problem with these random search methods is that the gene stability and genetic variability of the


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fragments selected this way are unknown. However, in our experiments the data obtained proved that our markers are stable and can be used for ecological studies. Introduced marker genes may affect the fitness of the engineered organisms. In addition, gene transfer to nonmarked organisms can not be excluded. (Look for details in later chapters of the book.) Therefore, intrinsic markers are generally considered to be more stable than introduced markers, but further data is needed from comparative studies to address this hypothesis. Intrinsic markers can be detected with remarkable sensitivity, as illustrated by the fact that detection of a single cell is possible from some samples. The performance of in situ hybridization with fluorescent probes and the results of some PCR-based detection techniques are comparable in this sense with those of engineered marker genes, but extensive optimization of assay conditions is required. An early criticism on nucleic acid-based detection techniques was based on their incapability to distinguish active and dead cells, but this problem can be overcome by the use of RNA-targeted probes. Detection of ribosomal RNA gives some insight into the dynamics of a bacterial population since number of ribosomes correlates with the metabolic activity of a cell. However, since rRNAs are extremely stable, they are not suitable targets for discriminating between living and nonliving cells. In contrast, detection of mRNA by hybridization or RT-PCR allows description of the active population relative to the total number of cells. Most of the molecular methods for detection of microorganisms based on intrinsic markers do not require isolation and cultivation of organisms, rather they allow in situ studies of individual members of complex microbial communities. By combined use of several probes with different specificities and by use of fingerprinting methods, different segments of a microbial population can be concurrently assessed. Therefore, the complementary use of several different methods will most likely be needed for accurate description of complex natural and man-made microbial communities.

Acknowledgments K. Lindström is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Cullen DW, Nicholson PS, Mendum TA et al. Monitoring genetically modified rhizobia in field soils using the polymerase chain reaction. J Appl Microbiol 1998; 84:1025-1034. 2. Olsen P, Wright S, Collins M et al. Patterns of reactivity between a panel of monoclonal antibodies and forage Rhizobium strains. Appl Environ Microbiol 1994; 60:654-661. 3. Faude UC, Höfle MG. Development and application of monoclonal antibodies for in situ detection of indigenous bacterial strains in aquatic ecosystems. Appl Environ Microbiol 1997; 63:4534-4542. 4. Trevors JT, van Elsas JD. A review of selected methods in environmental microbial genetics. Can J Microbiol 1989; 35:895-902. 5. Hames BD, Higgins SJ. Nucleic acid hybridization: a practical approach. Oxford.: IRL Press Ltd., 1985. 6. Saiki RK, Scharf S, Faloona F et al. Enzymatic amplification of β-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 1985; 230:1350-1354. 7. Saiki RK, Gelfand DH, Stoffel S et al. Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science 1988; 239:487-491. 8. Welsh J, McClelland M. Fingerprinting genomes using PCR with arbitrary primers. Nucleic Acids Res 1990; 24:7213-7218. 9. Williams JGK, Kubelik AR, Livak KJ et al. DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Res 1990; 22:6531-6535.


65

10. Lin J-J, Kuo J, Ma J. A PCR-based DNA fingerprinting technique: AFLP for molecular typing of bacteria. Nucleic Acids Res 1996; 24:3649-3650. 11. Martin B, Humbert O, Camara M et al. A highly conserved repeated DNA element located in the chromosome of Streptococcus pneumoniae. Nucleic Acids Res 1992; 20:3479-3483. 12. Versalovic J, Koeuth T, Lupski JR. Distribution of repetitive DNA sequences in eubacteria and application to fingerprinting of bacterial genomes. Nucleic Acids Res 1991; 24:6823-6831. 13. Muyzer G, de Waal EC, Uitterlinden AG. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 1993; 59:695-700. 14. Bej AK, Steffan RJ, DiCesare J et al. Detection of coliform bacteria in water by polymerase chain reaction and gene probes. Appl Environ Microbiol 1990; 56:307-314. 15. McDonald IR, Kenna EM, Murrell JC. Detection of methanotrophic bacteria in environmental samples with the PCR. Appl Environ Microbiol 1995; 61:116-121. 16. Cascón A, Anguita J, Hernanz C. Identification of Aeromonas hydrophila hybridization group 1 by PCR assays. Appl Environ Microbiol 1996; 62:1167-1170. 17. Shangkuan YH, Show YS, Wang TM. Multiplex polymerase chain reaction to detect toxigenic Vibrio cholerae O1. J Appl Bacteriol 1995; 79:264-273. 18. Hodson RE, Dustman WA, Garg RP et al. In situ PCR for visualization of microscale distribution of specific genes and gene products in procaryotic communities. Appl Environ Microbiol 1995; 61:4074-4082. 19. Ludwig W, Brockmann E, Beimfohr C et al. Nucleic acid based detection systems for genetically modified bacteria. System Appl Microbiol 1995; 18:477-485. 20. von Berg LK-H, Bothe H. The distribution of denitrifying bacteria in soils monitored by DNA-probing. FEMS Microbiol Ecol 1992; 86:331-340. 21. Brockman FJ. Nucleic-acid-based methods for monitoring the performance of in situ bioremediation. Mol Ecol 1995; 4:567-578. 22. Steffan RJ, Atlas RM. DNA amplification to enhance detection of genetically engineered bacteria in environmental samples. Appl Environ Microbiol 1988; 54:2185-2191. 23. Raaijmakers JM, Bitter W, Punte HLM et al. Siderophore receptor PupA as a marker to monitor wild-type Pseudomonas putida WCS358 in natural environments. Appl Environ Microbiol 1994; 60:1184-1190. 24. Jacobsen CS. Microscale detection of specific bacterial DNA in soil with a magnetic capture-hybridization and PCR amplification assay. Appl Environ Microbiol 1995; 61: 3347-3352. 25. Jansson JK. Tracking genetically engineered microorganisms in nature. Curr Opin Biotechnol 1995; 6:275-283. 26. Hwang I, Farrand SK. A novel gene tag for identifying microorganisms released into the environment. Appl Environ Microbiol 1994; 60:913-920. 27. Flemming CA, Leung KT, Lee H et al. Survival of lux-lac-marked biosurfactant-producing Pseudomonas aeruginosa UG2L in soil monitored by nonselective plating and PCR. Appl Environ Microbiol 1994; 60:1606-1613. 28. Van Elsas JD, Van Overbeek LS, Fouchier R. A specific marker, pat, for studying the fate of introduced bacteria and their DNA in soil using a combination of detection techniques. Plant Soil 1991; 138:49-60. 29. Möller A, Gustafsson K, Jansson JK. Specific monitoring by PCR amplification and bioluminescence of firefly luciferase gene-tagged bacteria added to environmental samples. FEMS Microbiol Ecol 1994; 15:193-206. 30. Pillai SD, Pepper IL. Transposon Tn5 as an identifiable marker in rhizobia: survival and genetic stability of Tn5 mutant bean rhizobia under temperature stressed conditions in desert soils. Microb Ecol 1991; 21:21-33. 31. van Elsas JD, Nikkel M, van Overbeek LS. Detection of plasmid RP4 transfer in soil and rhizosphere, and the occurrence of homology to RP4 in soil bacteria. Current Microbiology 1989; 19:375-381.

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32. Gürtler V, Stanisich VA. New approaches to typing and identification of bacteria using the 16S-23S rDNA spacer region. Microbiology 1996; 142:3-16. 33. Amann RI, Ludwig W, Schleifer K-H. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Rev 1995; 59:143-169. 34. Ward DM, Bateson MM, Weller R et al. Ribosomal RNA analysis of microorganisms as they occur in nature. In: Marshall KC, editor. Advances in Microbial Ecology. v. 12. New York: Plenum Press, 1992:219-86. 35. Amann RI. Fluorescently labelled, rRNA-targeted oligonucleotide probes in the study of microbial ecology. Mol Ecol 1995; 4:543-554. 36. Devereux R, Kane MD, Winfrey J et al. Genus- and group-specific hybridization probes for determinative and environmental studies of sulfate-reducing bacteria. System Appl Microbiol 1992; 15:601-609. 37. Briglia M, Eggen RIL, De Vos WM et al. Rapid and sensitive method for the detection of Mycobacterium chlorophenolicum PCP-1 in soil based on 16S rRNA gene-targeted PCR. Appl Environ Microbiol 1996; 62:1478-1480. 38. Young JPW. Phylogeny and taxonomy of rhizobia. Plant Soil 1996; 186:45-52. 39. Olsen GJ, Woese CR. Ribosomal RNA: a key to phylogeny. FASEB J 1993; 7:113-123. 40. Clayton RA, Sutton G, Hinkle PS et al. Intraspecific variation in small-subunit rRNA sequences in GenBank: Why single sequences may not adequately represent prokaryotic taxa. Int J Syst Bacteriol 1995; 45:595-599. 41. Schmidhuber S, Ludwig W, Schleifer KH. Construction of a DNA probe for the specific identification of Streptococcus oralis. J Clin Microbiol 1988; 26:1042-1044. 42. Liang P, Pardee AB. Differential display of eukaryotic messenger RNA by means of the polymerase chain reaction. Science 1992; 257:967-971. 43. Triplett EW, Sadowsky MJ. Genetics of competition for nodulation of legumes. Annu Rev Microbiol 1992; 46:399-428. 44. Paau AS. Improvement of Rhizobium inoculants by mutation, genetic engineering and formulation. Biotech Adv 1991; 9:173-184. 45. Lindström K. Rhizobium galegae, a new species of legume root nodule bacteria. Int J Syst Bacteriol 1989; 39:365-367. 46. Lipsanen P, Lindström K. Infection and root nodule structure in the Rhizobium galegae sp. nov.-Galega symbiosis. Symbiosis 1988; 6:81-96. 47. Lindström K, Jurgens G, Kaijalainen S et al. Rhizobium galegae - properties, phylogeny and molecular identification methods. In: Kiss GB, Endre G, editors. Proceedings of the 1st european nitrogen fixation conference and the workshop “Safe application of genetically modified microorganisms in the environment”. Szeged, Hungary: Officina press, 1994:265-9. 48. Lindström K, Lipsanen P, Kaijalainen S. Stability of markers used for identification of two Rhizobium galegae inoculant strains after five years in the field. Appl Environ Microbiol 1990; 56:444-450. 49. Suominen L, Saano A, Lindström K. Cloning of the host specific gene region of Rhizobium galegae. In: Gresshoff PM, Roth LE, Stacey G, et al., editors. Nitrogen fixation: achievements and objectives. New York: Chapman and Hall, 1990:589. 50. Saano A, Kaijalainen S, Lindström K. Isolation of Rhizobium galegae species-specific DNA probes. In: Gresshoff PM, Roth LE, Stacey G, et al., editors. Nitrogen fixation: achievements and objectives. New York: Chapman and Hall, 1990:575. 51. Tas É, Leinonen P, Saano A et al. Assessment of competitiveness of rhizobia infecting Galega orientalis on the basis of plant yield, nodulation and strain identification by antibiotic resistance and PCR. Appl Environ Microbiol 1996; 62:529-535. 52. Welcher AA, Torres AR, Ward DC. Selective enrichment of specific DNA, cDNA and RNA sequences using biotinylated probes, avidin and copper-chelate agarose. Nucleic Acids Res 1986; 14:10027-10044. 53. Straus D, Ausubel F. Genomic subtraction for cloning DNA corresponding to deletion mutations. Proc Natl Acad Sci USA 1990; 87:1889-1893.


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54. Bjourson AJ, Cooper JE. Isolation of Rhizobium loti strain-specific DNA sequences by subtraction hybridization. Appl Environ Microbiol 1988; 54:2852-2855. 55. Streit W, Bjourson AJ, Cooper JE et al. Application of subtraction hybridization for the development of Rhizobium leguminosarum biovar phaseoli and Rhizobium tropici groupspecific DNA probe. FEMS Microbiol Ecol 1993; 13:59-68. 56. Sargent TD, Dawid IB. Differential gene expression in the gastrula of Xenopus levis. Science 1983; 222:135-139. 57. Kunkel LM, Monaco AP, Middleworth W et al. Specific cloning of DNA fragments absent from the DNA of a male patient with an X chromosome deletion. Proc Natl Acad Sci USA 1985; 82:4778-4782. 58. Bjourson AJ, Stone CE, Cooper JE. Combined subtraction hybridization and polymerase chain reaction amplification procedure for isolation of strain-specific Rhizobium DNA sequences. Appl Environ Microbiol 1992; 58:2296-2301. 59. Tas É, Kaijalainen S, Saano A et al. Isolation of a Rhizobium galegae strain-specific DNA probe. Microb Releases 1994; 2:231-237. 60. Darrasse A, Kotoujansky A, Bertheau Y. Isolation by genomic subtraction of DNA probes specific for Erwinia carotovora subsp. atroseptica. Appl Environ Microbiol 1994; 60:298-306. 61. Tas É, Saano A, Leinonen P et al. Identification of Rhizobium spp. in peat-based inoculants by DNA hybridization and PCR and its application in inoculant quality control. Appl Environ Microbiol 1995; 61:1822-1827. 62. Nick G, Lindström K. Use of repetitive sequences and the polymerase chain reaction to fingerprint the genomic DNA of Rhizobium galegae strains and to identify the DNA obtained by sonicating the liquid cultures and root nodules. System Appl Microbiol 1994; 17:265-273. 63. Alm EW, Oerther DB, Larsen N et al. The oligonucleotide probe database. Appl Environ Microbiol 1996; 62:3557-3559. 64. Maidak BL, Cole JR, Parker CTJ et al. A new version of the RDP (Ribosomal Database Project). Nucleic Acids Res 1999; 27:171-173. 65. Gottschal JC, Meijer WG, Oda Y. Use of molecular probing to assess microbial activities in natural ecosystems. In: Insam H, Rangger A, editors. Microbial communities. Berlin: Springer-Verlag, 1997:10-8. 66. Klein PG, Juneja VK. Sensitive detection of viable Listeria monocytogenes by reverse transcription-PCR. Appl Environ Microbiol 1997; 63:4441-4448. 67. Felske A, Akkermans ADL, De Vos WM. Quantification of 16S rRNAs in complex bacterial communities by multiple competitive reverse transcription.PCR in temperature gradient gel electrophoresis fingerprints. Appl Environ Microbiol 1998; 64:4581-4587. 68. Eichner CA, Erb RW, Timmis KN et al. Thermal gradient gel electrophoresis analysis of bioprotection from pollutant schocks in the activated sludge microbial community. Appl Environ Microbiol 1999; 65:102-109. 69. Cheng J, Sheldon EL, Wu L et al. Preparation and hybridization analysis of DNA/RNA from E. coli on microfabricated bioelectronic chips. Nature Biotechnology 1998; 16:541-546. 70. Drolet DW, Moon-McDermott L, Romig TS. An enzyme-linked oligonucleotide assay. Nature Biotechnology 1996; 14:1021-1025. 71. Egholm M, Buchardt O, Christensen L et al. PNA hybridizes to complementary oligonucleotides obeying the Watson-Crick hydrogen-bonding rules. Nature 1993; 365:566-568. 72. Tuerk C, Gold L. Systematic evolution of ligands by exponential enrichment: RNA ligands to bacteriophage T4 DNA polymerase. Science 1990; 249:505-510. 73. Gold L. Oligonucleotides as research, diagnostic and therapeutic agents. J Biol Chem 1995; 270:13581-13584. 74. Langworthy DE, Stapleton RD, Sayler G et al. Genotypic and phenotypic response of a riverine microbial community to polycyclic aromatic hydrocarbon contamination. Appl Environ Microbiol 1998; 64:3422-3428.

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75. Pennanen T, Fritze H, Vanhala P et al. Structure of a microbial community in soil after prolonged addition of low levels of simulated acid rain. Appl Environ Microbiol 1998; 64:2173-2180.

CHAPTER 5

Luminescence-Based Microbial Marker Systems and Their Application in Microbial Ecology James I. Prosser, Antonio J. Palomares, Matti T. Karp and Philip J. Hill

5.1. Introduction

M

olecular marker systems provide the ability to track specific microbial inocula in the environment. This is a fundamental requirement for many ecological studies and is particularly appropriate for the assessment of risks associated with the release of genetically modified microorganisms. The efficiency of a marker system requires the absence of the marker gene, or its phenotype, from the environment under study, stable insertion of the marker genes into the host organism without loss of fitness and efficient and sensitive genotypic and/or phenotypic detection. Techniques for genotypic detection are similar for all marker genes and distinctions therefore arise through differences in efficiency of phenotype detection and the information that this provides. The major advantage of all marker systems over traditional techniques is the increased ability to distinguish specific organisms under study from the indigenous population. Marker systems do, however, require laboratory cultivation of the host strain and ecological studies utilizing marked strains must accept this limitation. This is not, however, a limitation for applications to risk assessment of genetically modified microorganisms, which require growth in the laboratory before release. This article focuses on luminescence-based techniques that involve marking with prokaryotic or eukaryotic genes encoding luciferase, which catalyses light production. They, like all marker systems, increase selective detection of inocula but provide several additional advantages. In particular, they enable rapid and specific measurement of metabolic activity of marked organisms, without cell extraction, and studies of their spatial distribution in environmental samples.

5.2. Lux-Based Systems Bacterial luciferase catalyses the oxidation of FMNH2 and a long chain aldehyde by molecular oxygen to produce FMN and the corresponding acid, with concomitant emission of blue-green light:1 RCHO + O2 + FMNH2 → RCOOH + FMN + H2O + hν (ca. 490 nm in Vibrio spp.) Luciferase is a heterodimeric enzyme of ca. 79 kDa that is cytoplasmic in Vibrio harveyi and membrane-bound in Plesiomonas leiognathi. The reaction2 is initiated by an interaction between FMNH2 and luciferase to produce luciferase-bound FMNH2 (Intermediate I), which Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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reacts with molecular oxygen to form 4α-hydroperoxy-FMNH (Intermediate II) (Fig. 5.1). In the absence of aldehyde, this decays to generate FMN and hydrogen peroxide with no light emission (dark decay). In the presence of aldehyde, Intermediate III (4α peroxyhemiacetal-FMNH) is formed, which decays to yield the aliphatic acid and an excited emitter, proposed to be a 4α-hydroxy-FMNH species,3 which undergoes a radiative relaxation to emit light. After returning to the ground state, 4α-hydroxy-FMNH decays to generate water and oxidized FMN. Although the in vivo aldehyde substrate appears to be tetradecanal, other long chain aldehydes (e.g., nonanal or decanal) may elicit a bioluminescent response. Enzyme turnover rate is greatly affected by the nature of the aldehyde substrate used, and may differ significantly and characteristically with luciferases from different species.

5.2.1. Genetic Organization of the lux Regulon The genes encoding bacterial luciferase and the other proteins required for bioluminescence are situated in lux operons, which have been most studied in Photobacterium and Vibrio. In Vibrio fischeri, all lux genes can be assigned to two operons, L and R, whose organization is illustrated in Figure 5.2.4 The only known gene in operon L is luxR, which encodes a regulatory protein. Operon R contains luxA and luxB, which code for the α and β subunits of luciferase respectively, and luxC, D, E and I, which respectively code for a fatty acid reductase, an acyl transferase, an acylprotein synthase and synthesis of the ‘autoinducer’, N-(3-oxo-hexanoyl) homoserine lactone, which induces light production in all V. fischeri strains. In V. fischeri strains MJ-1 and ATCC7744 these genes are in the order luxICDABE. The organization of structural genes for bioluminescence in other luminescent bacteria is similar, except that regulatory genes are omitted.5 Another gene, luxF, lying between luxB and luxE and sharing high homology with luxB, has been identified in Photobacterium phosphoreum and P. leiognathi.6,7 An additional gene, luxG, has been identified downstream of luxE in all marine bioluminescent bacteria, but not in any Photorhabdus luminescens

Fig. 5.1. The mechanism of bacterial bioluminescence. L represents luciferase, roman numerals show key intermediates, * indicates excited intermediate.

Luminescence-Based Microbial Marker Systems and Their Application in Microbial Ecology 71

Fig. 5.2. A physical map of the lux regulon.

strains studied,5 and gene luxH has been identified downstream of luxG in V. harveyi.8 These genes are not essential for a bioluminescent phenotype but may be involved in the generation of reduced flavin substrate for the luminescence reaction. All luxA and luxB genes sequenced (e.g., refs. 7,9) show a high degree of sequence homology. The luxA gene shows greater conservation than luxB, consistent with the postulate that the α subunit controls the kinetic properties of the enzyme. Sequence analysis indicates that bacterial luciferases may be subdivided into two distinct classes, the first represented by those from V. fischeri and Photobacterium and the second by V. harveyi, P. luminescens and a symbiont from the fish Kryptophanaron alfredi, the latter group comprising the less heat-labile enzymes. The high homology between α and β subunits both within and between different species indicates that luxA and luxB arose by gene duplication.5 The α subunit appears to be solely responsible for the catalytic activity of the enzyme but the β subunit is essential for functionality, and may be required to maintain the catalytically active conformation of α residues.

5.2.2. Supply of Aldehyde for the Bioluminescent Reaction The bioluminescent reaction requires a long chain aldehyde substrate and tetradecanoic acid appears to be the precursor of the natural substrate tetradecanal.10 A fatty acid reductase complex, responsible for aldehyde synthesis, consists of two distinct functional components;11 acyl protein synthase (50 kDa) which, with cleavage of ATP to AMP, activates the fatty acid to a fatty acyl intermediate, and acyl-protein reductase (58 kDa), responsible for the transfer of the acyl group from the synthase and its subsequent NADPH dependent reduction to release free tetradecanal. A third component, initially identified in P. phosphoreum,12 is a 34 kDa acyl transferase and is acylated only in the presence of the other two polypeptides. It utilizes acyl-acyl carrier protein as substrate to supply free fatty acid to the luminescent system13 but is unnecessary for fatty acid reductase activity in vitro. Aldehyde synthesis involves channeling of tetradecanoic acid from acyl-ACP, by the acyl protein transferase subunit, releasing the free acid to acylate the acyl protein synthase in an ATP dependent reaction. After removal of AMP, the acyl group is transferred in a freely reversible reaction to the acyl protein reductase subunit, which, utilizing the reducing power of NADPH, releases free tetradecanal. This aldehyde is oxidized by luciferase, with concomitant light emission, back to the fatty acid that may then be recycled.

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5.2.3. Expression Vectors for Bacterial Luciferases Cloning vectors using insertional inactivation of luxAB as a marker gene, instead of the ubiquitous LacZα complementation, have been reported, but most published uses of luciferase encoding plasmid vectors concern their use in the study of DNA control sequences. For example, the construction of vectors containing promoters upstream of luxAB with a polylinker between the promoter and lux genes allows for the identification of transcriptional terminators as evidenced by decreased bioluminescence of clones containing terminators.14 Many luxAB containing plasmids have been employed as promoter probe vectors, allowing detection and determination of strength of promoter DNA cloned upstream of luxAB or a whole lux operon. Many such vectors are available for use in gram-negative and grampositive bacteria. Another method of promoter probing is to use transposons. In addition to providing insertional inactivation of chromosomal genes, lux-containing transposons may allow the expression of target genes to be monitored on the basis of bioluminescence. A number of such transposons have been reported, including a miniMu-lux in E. coli and other gram-negative bacteria,15 Tn5 derivatives containing luxAB in Pseudomonas16 and other bacteria and a Tn1721 derivative in X. campestris.17 These transposon systems have been used successfully to identify environmentally responsive genes,18 maintain a constitutive expression of bioluminescence in normally dark bacteria,19 for promoter probing16 and in the characterization of loci for phytopathogenicity.17 More recently, mini-Tn5 derivatives containing the whole lux operon from P. luminescens, obviating the need to add exogenous aldehyde, have been constructed and shown to be effective in many gram-negative organisms.20 Shuttle plasmid vectors, constructed for expression of either bacterial or insect luciferase genes in grampositive and gram-negative strains, have been found to lead to high levels of light emission which could be measured in real-time from intact cells.21

5.3. Luc-Based Systems Although bacterial luciferases are most frequently used as markers for environmental use, strains carrying the firefly luciferase or other insect luciferase genes can provide advantages. The enzymes from insects are in general more heat stable, are not subject to substrate inhibition and their quantum yield (90%) is considerably higher than that of bacterial luciferase (5-10%). Firefly luciferase requires luciferin, a heterocyclic carboxylic acid, oxygen and ATP as substrates. The requirement for ATP is very specific and firefly luciferase cannot utilize other nucleotide triphosphates. In the presence of magnesium and molecular oxygen, luciferase catalyses the oxidative decarboxylation of luciferin, resulting in the production of light and oxyluciferin, with hydrolysis of ATP. D-Luciferin + ATP + Mg → Luciferase-luciferyl-AMP + PPi Luciferase-luciferyl-AMP + O2 → oxyluciferin + AMP+CO2 + light This reaction has a quantum yield of 0.88 photons emitted per luciferin oxidized, and is the most efficient luminescent reaction known.22 The best-characterized firefly luciferase is that of the common North American firefly Photinus pyralis, from which luciferase has been purified and the luciferin substrate determined.23 SDS-polyacrylamide gel electrophoresis of purified P. pyralis luciferase shows a single band of protein with an apparent molecular weight of 62,000.24 Only one dehydroluciferyl adenylate, a substrate intermediate analogue, binds per luciferase dimer although there are two ATP and two luciferin binding sites.25 This suggests that the two subunits in the dimer may be different, but it has not been possible to separate physically two forms of the protein.


A potential disadvantage of luc systems is the requirement for exogenous addition of luciferin. However, uptake of D-luciferin is efficient at slightly acidic conditions, e.g., pH values of 5 and 6 for E. coli26 and Streptococcus mutans,27 respectively. A thermostable variant of firefly luciferase has also been generated by genetic engineering28 and is able to sustain temperatures up to 50˚C for 60 minutes, in vitro. The color of light emitted by firefly luciferase varies between species. P. pyralis emits yellow-green light, with the peak emission occurring at 562 nm. This color can be shifted to red in vitro by adding divalent metal cations, such as zinc, by increasing the temperature, or by reducing the pH to below 7.0.22 Light emitted by other species of firefly ranges from green (550 nm) to yellow (579 nm), possibly reflecting adaptation to the time of night during which fireflies are active. Twilight-active species tend to emit yellow light that should be more visible against the background light than the green light of dark-active species. The Jamaican click beetle, Pyrophorus plagiophthalamus, has a pair of dorsal organs that emit bright green light (543 nm) and a ventral organ that emits yellow-orange light (582 nm).22 The color of light emitted is a property of the luciferase, rather than the luciferase substrate or filtering of the light by pigments.22 The ability of all beetle luciferases to utilize synthetic P. pyralis luciferin suggests that all synthesize very similar luciferin, and that differences in the color of the emitted light must be due to structural differences in the luciferases themselves. Furthermore, there must be extensive conservation in the overall structure of the beetle luciferases since they must all possess the same substrate binding sites. This is supported by the observation that anti-P. pyralis luciferase antibodies recognize luciferases from the six species of firefly tested as well as the luciferase from the click beetle P. plagiophthalamus. Fireflies have long life cycles and are not easily maintained in the laboratory, making genetic approaches to the study of firefly luciferase impractical. The cloning and expression of firefly luciferase genes in heterologous systems, however, enables the creation of mutant luciferases in vitro by means of recombination technology.

5.3.1. Expression Vectors Using Eukaryotic Luciferases as Bacterial Markers The eukaryotic nature and presumed absence of firefly and click beetle luciferase genes may provide a unique genotype to bacteria and the variation in wavelength of emitted light provides an additional advantage. To exploit this difference in luciferase assays, luc and lucOR genes, which encode emission of light at 560 nm (yellow-green) and 595 nm (orange), respectively, have been used to develop marker genes for bacteria employing three transcriptional units (Fig. 5.3). The first, an antitetracycline PI promoter, provides considerable constitutive expression when P1::luc and P1::lucOR are borne on ColE1 plasmids, but luciferase activity decreases when both fusions are borne on a RK2 derivative plasmid. This reduction is due to the presence of a tetR gene in the parental plasmid pRK293 or to differences in copy number. A second system, cI857-λPR, provides a highly regulated promoter which is useful for marking cells for environmental release, as it may be silenced in the natural environment, avoiding the use of nutrients or energy in marker synthesis. In the presence of the cI857 repressor gene, λPR is a well-regulated promoter,29 as found when the PI::luc fusion is expressed in E. coli in either ColE1 or RK2 replicons. Detection of marked organisms requires induction of the marker gene, allowing identification on solid media or in enrichment broth. In the presence of the cI857 gene, λPR is not efficiently regulated in some gram-negative bacteria but constitutive expression of either luc or lucOR from λPR can be achieved efficiently in a range of bacteria. 30 λPR expresses high levels of luciferase activity, which are easily detected constitutively. The third system involves a strong promoter, Ptrc, which provides higher and better-regulated luminescence in bacteria that express lucOR and leads to greater in vivo luciferase activity in E. coli and other gram-negative bacteria. Ptrc is effective in a range of bacterial hosts and, following induction, yields

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Fig. 5.3. Plasmids constructed to study the expression of eukaryotic luciferases under the control of different promoters in gram-negative bacteria. All of the plasmids were based on the RK2 derivative pRK293. Fragments with the represented fusions of luc and lucOR were cloned in the sites of pRK293 indicated. B, BamHI; Bg, BglII; C, ClaI; H, HindIII S, SalI; X, XhoI. Reprinted with permission from Cebolla A, Vázquez ME, Palomares AJ. Appl Environ Microbiol 1995; 61:660-668.

higher levels of luminescence than other constructs.31 Use of λPR::luc and λP R::lucOR enables distinction between Pseudomonas putida colonies emitting light of different wavelengths30 (Fig. 5.4).

5.4. Methodology Luminescence-marked organisms can be monitored using several techniques. Lux and luc genes may be detected by gene probing, but the major advantages of luminescence marker systems lie in the ability to detect light.

5.4.1. Detection of Luminescence by Eye and by Photographic and X-ray Film Viable cell enumeration of luciferase-marked strains can be achieved using traditional dilution plate counting and luminescent colonies can usually be detected by the unaided eye in a darkened room.32 Greater sensitivity can be achieved using photographic film (with long exposure), X-ray film or photon imaging (see below). The majority of luciferase-marked strains are also marked with antibiotic resistance (see Chapter 2), enabling selectivity against indigenous populations by enumeration on media supplemented with the appropriate antibiotic. In the absence of antibiotic resistance markers, selective enumeration depends on the competitive ability of the marked strain on enumeration media, the nutrients required for luminescence and the sensitivity of detection. Nevertheless, visual detection of a single luminescent colony of lux-marked Erwinia carotovora is possible in the presence of 3000 nonluminescent colonies.32 Similar techniques may be used for in situ detection of luminescent strains. Photography has also been used to compare in vivo activities of firefly and click


Fig. 5.4. Luminescence emitted by two strains of Pseudomonas putida containing plasmids pAP2 and pACR3 with PR::luc and PR::lucOR fusions, respectively. Cells were incubated on nitrocellulose filters placed on solid medium, incubated for 30 h and photographed under normal illumination (A) and after 30 min exposure in the dark. Reprinted with permission from Cebolla A, Vázquez ME, Palomares AJ. Appl Environ Microbiol 1995; 61:660-668.

beetle luciferase in E. coli cultures on nitrocellulose filters, with exposure for 10-15 minutes.30 Similar examination of E. coli streaks on solid media demonstrated the four colors of bioluminescence emitted by different click beetle luciferases, with 2 s exposure time.33 This approach has also been used to determine regions of plants infected with lux-marked pathogenic bacteria 34 and for detection of luciferase activity in alfalfa nodules induced by R. meliloti containing a nifH::luc fusion.35

5.4.2. Luminometry The major advantage of lux- and luc- marker systems lies in the relationship between luminescence and cell activity enabling assessment of the activity of marked organisms by quantification of luminescence, by luminometry. In organisms in which luciferase production is constitutive and where luminescence per cell is maximal, or constant, total light emission is proportional to biomass concentration. This occurs in exponentially growing cultures, with proportionality between luminescence and biomass concentration over several orders of magnitude.27,36 The convenience and validity of this approach depend on the nature of the construct. For example, constitutive luciferase production may be achieved by chromosomal marking with luxA and luxB genes, but luminescence requires addition of dodecanal and incubation under optimal conditions. Cellular production of dodecanal is achieved in cells additionally marked with luxCDE genes, but the additional genetic load may affect the physiological characteristics of the marked organism. Increased luminescence may also be achieved by increasing lux-or luc- gene copy number, for example, in plasmid marked strains, although this increases the likelihood of decreased host fitness and stability.37 Finally, use of luminometry in samples with particulate material reduces sensitivity. Lower detection limits using luminome try therefore depend on a number of factors. For example, detection limits for chromosomally and plasmid-marked strains of P. fluorescens in cell suspensions were 1.7 x 103 and 8.9 x 104 cells ml-1, respectively. Equivalent detection limits in inoculated autoclaved sterile soil were 8.1 x 103 and 3.0 x 105 cell g-1, respectively,

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while inoculation into nonsterile soil increased detection limits to 5.9 x 104 and 2.2 x 105 cells g-1, possibly through greater competition for nutrients.38 An added advantage of chromosomal marking is the frequent lack of apparent effect on host fitness. In the environment, conditions are likely to be variable and sub-optimal for luminescence. Luminometry of untreated samples will therefore provide a measure of the activity of marked organisms under nutritional and environmental conditions prevailing at the time of sampling. Luminescence activity is equivalent to that measured using traditional techniques, e.g., dehydrogenase activity39 but luminometric measurements are considerably more sensitive, convenient and rapid, with results within several minutes of sampling. More importantly, luminometry assesses the activity of the marked population only, in the presence of high indigenous populations, while traditional activity techniques, e.g., enzyme assays, substrate conversion rates, provide information on the total population. The technique is also sufficiently sensitive to detect the activity of viable but nonculturable populations.40 Luminescence assays may be modified to assess biomass by amendment with nutrients and incubation under optimal conditions.41 This measure of potential luminescence is equivalent to other measures of potential activity, e.g., the substrate induced respiration method.

5.4.3.Photon Imaging Quantitative imaging of gene expression using luciferase based reporter constructs is now possible due to the availability of extremely sensitive imaging equipment that can be calibrated using light standards. These camera systems are usually based on image intensifiers coupled to video cameras, or cooled charge coupled device (CCD) technology. Intensified cameras have high sensitivity and the ability to watch the image form almost in realtime but, at high gain, suffer from reduction in spatial resolution. Cooled CCD systems rely on capturing the image during a single long exposure. They have extremely low noise and wide dynamic range and are increasingly available, leading to a reduction in cost of this technology. The sensitivity of many cooled CCD cameras can be significantly increased by pixel binning, although this reduces spatial resolution. Both systems may be configured for microscopic analysis or imaging of macro samples. They are often supplied with flexible image processing software that allows the user to count areas, quantify signals and superimpose images. Examples of the power of this technology to assess gene expression, even in highly complex environments, include assessment of mas gene (mannopine synthesis) expression by Agrobacterium tumefaciens in different tobacco plant tissues during infection42 and the coupling of luxAB with the nitrogenase promoters of nifD and nifH from Bradyrhizobium japonicum, enabling imaging of luciferase expression in single soybean root nodules and, indeed, in single bacteroid infected cells.43

5.5. Applications 5.5.1. Stable Tagging of Bacteria with Firefly Luciferase Gene for Environmental Monitoring Gram-negative strains engineered for in situ applications must meet a number of practical requirements for safe and efficient performance. These include not only the absence of traits that may give them an advantage over nonengineered strains but also the ability to retain the acquired genotype and phenotype in the absence of selection. Use of firefly luciferase addresses the first of these concerns, since D-luciferin is expected to be absent from all prokaryotic cells and the enzyme is unlikely to provide any advantage over the wild type. Stable inheritance of the marker gene, in the absence of selective pressure, can be achieved by chromosomal marking using the mini-Tn5 system.44 A HindIII fragment containing only


the λPR promoter and the luc genes was isolated and cloned into pUC18Not. The NotI fragment with the luc gene was cloned into the unique NotI site of pUT/mini-Tn5 Sm/Sp and into pUTKm. The resulting plasmids, pTCR210 and pTCR240, were selected for studies of transposition and luciferase expression in various gram-negative bacteria. 36 To tag bacteria without the use of antibiotic resistance genes, the Km resistance gene may be replaced with the lPR::luc fusion. To attenuate the expression in the environment, where the marked strain must survive, and to allow sensitive detection, a DNA fragment containing the repressor gene lacIq and a PR::lucOR fusion has been introduced on a suicide plasmid. PTEB510 (Fig. 5.5) is a representative of constructs able to express high luciferase levels only when induced by IPTG. Transposition frequencies were suitable for marking purposes and the levels of luminescence were sufficient for detection by the standard methods.

Fig. 5.5. Construction of suicide orange click beetle luciferase plasmids used for marking gramnegative bacteria. The selective markers are flanked by Tn5 19-bp terminal ends (I end and O end). The IS50R tnp gene devoid of NotI sites (tnp*) is oriented divergently from the I end. Restriction sites: E, EcoR1; Nt, NotI; Xb, XbaI; H, HindIII; B, BamHI; S, SalI; P, PstI; Bg, BglII. Reprinted with permission from Vázquez ME, Cebolla A, Palomares, AJ. FEMS Microbiol Lett 1994; 121:11-18.

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5.5.2. Determination of Rhizobium Nodule Occupancy by Luminescence The symbiotic interaction between rhizobia and leguminous plants results in the formation of root nodules through a number of stages determined by different sets of genes both in the microsymbiont and macrosymbiont. Quantification of nodule occupancy is required to determine the competitiveness of two or more strains in producing and occupying nodules. Luminescence marking of bacteria that interact with plants provides a simple and highly sensitive system for their detection, particularly under conditions in which light emission is directly related to cell concentration. Palomares et al.45 compared luc-marked R. meliloti strain CR201 with the wild-type strain, R. meliloti 1021. To determine the competitiveness of the marked luminescent strain, alfalfa plants were inoculated with suspensions containing known proportions of each strain. Luminescent and nonluminescent cells isolated from nodules were enumerated by plate counting and the proportions of the two strains were equal to those in the inocula. Under the experimental conditions employed, light emission was linearly related to luciferase activity over at least four orders of magnitude.46 Luciferase activity was then determined in bacteroid suspensions taken from nodules inoculated with different proportions of luminescence-marked cells and was found to be directly related to viable cell concentrations. Direct measurement of luminescence of cells isolated from nodules provides many advantages over other methods. In particular, it does not require plating and is less laborious and less time consuming than alternative marker phenotypes that require colony formation.

5.5.3. Eukaryotic Luciferase Markers for Assessment of Plasmid Transfer in Soil Environments Most studies of gene transfer in the soil have focused on conjugal transfer and there are concerns regarding the transfer of the genetically engineered DNA sequences to indigenous microorganisms. Vázquez and Palomares46 have established a model system for plasmid transfer analysis involving constructs obtained using the lambda promoter, λPR, and the luciferase genes luc and lucOR (see above). This promoter is regulated by the thermosensitive repressor cI857 that is active at 28˚C but not at 42˚C, at which temperature the promoter, λPR, is induced. A luciferase fusion system was constructed consisting of cI857-λPR::luc and cI857λPR::lucOR which is temperature-inducible in E. coli. Expression was constitutive at 28˚C and luminescence could be detected using either luc or lucOR genes when the plasmids were mobilized to other backgrounds. Use of luciferase genes encoding emission of either green or orange light demonstrated transfer from RK2 derivative plasmids, incompatibility group IncP (pACR4 and pACR18), and from R388 derivative plasmids, incompatibility group IncW (pET2), either in vitro or in microcosms of sterile and nonsterile soil. Mobilization from RK2 and R388 derivatives to P. putida 2440 (co-introduced with donors) or indigenous soil organisms was demonstrated. High transfer frequencies were found for both plasmid groups in assays on solid media. The experiments in sterile soil enabled assessment of the influence of abiotic environmental parameters on gene transfer; in this case transfer frequency was two orders of magnitude lower than on solid media, perhaps due to physical separation between donor and receptor.

5.5.4. Survival and Resuscitation of Microbial Inocula Luminescence-based markers provide the ideal strategy for risk assessment of genetically engineered microorganisms following their release into the environment. This requires monitoring of population size and activity to quantify survival, growth and recovery following periods of stress or starvation. An example of this approach is assessment of the effect of matric potential on the survival of a strain of Pseudomonas fluorescens, 10586s/ FAC510, chromosomally marked with lux genes and kanamycin resistance.47 This strain


was inoculated into microcosms containing autoclaved or nonautoclaved soil after adjustment of matric potential to -30, -750 or -1500 kPa. Selective viable cell enumeration was achieved by plate counting on media containing kanamycin and population activity was measured by luminometry, radiorespirometry and dehydrogenase activity. Matric potential did not significantly affect survival, as determined by plate counts, but survival was significantly greater in autoclaved soil at higher matric potentials. This may be due to reduced competition and predation, as protozoan movement will be lower at the lowest matric potential. Luminescence activity decreased rapidly following inoculation, and at a greater rate than viable cell counts, and was the most sensitive indicator of effects of matric potential and autoclaving. It was also found to be significantly more sensitive, reproducible and convenient as a measure of population activity than traditional techniques of radiorespirometry and dehydrogenase activity in autoclaved soil. More importantly, in nonautoclaved soil, luminescence provided a measure of the marked organism only in the presence of a significant indigenous microbial population. Potential luminescence involved assessment of luminescence during incubation with nutrients and indicated the speed of recovery of starved organisms. Early samples showed no increase in luminescence during the incubation period at the highest matric potentials, indicating little inactivation immediately following inoculation, but significant increases were observed during incubation of samples from soil at the lowest matric potential. The rate of recovery and final potential luminescence values decreased with time spent by cells in the soil and with decreasing matric potential, and were lowest in nonautoclaved soil. Samples taken from nonautoclaved soil at the lowest matric potential after 28 days showed no recovery. This suggests that P. fluorescens, inoculated into soil at this matric potential, would have negligible environmental impact within four weeks of inoculation, as any available substrates would be rapidly utilized by competing indigenous organisms. Conversely, the lack of persistence of activity of this organism might question its suitability in performing the function for which it was originally designed. A combination of viable cell enumeration, luminescence and potential luminescence measurements therefore provides information which is of relevance and importance to both risk assessment and the development of more efficient microbial inocula.

5.5.5. Predation The ability to detect luminescence activity of specific marked strains has been exploited to determine the effect of predation by protozoa on bacterial prey.48 Soil microcosms were inoculated with lux-marked bacterial cells (P. fluorescens) at specific matric potentials to ensure their location in small (neck size < 6 µm) or intermediate size (neck size 6-30 µm) pores, while Colpoda steinii was introduced to large pores (neck size 30-60 µm) (Fig. 5.6). Viable cell concentrations of both populations were monitored and bacterial activity was assessed by luminometry. Bacteria in intermediate pores were predated at a greater rate than those introduced to small pores, and decreases in bacterial viable cell concentration were associated with increases in the ciliate population. Small pores therefore appear to provide protection from predation. Luminescence decreased, following inoculation, due to loss of bacterial activity through lack of nutrients and this effect was greater in the presence of the ciliate, due to predation. However, luminescence activity per cell was significantly greater for cells introduced into intermediate-sized pores, where predation was greatest. This suggests that predation, although reducing numbers of bacteria, results in the release and turnover of nutrients by the ciliate such that surviving bacterial cells have greater activity. Demonstration of this effect using traditional activity techniques was not possible because of problems in distinguishing between bacterial and ciliate activity.

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Fig. 5.6. Changes in a) viable cell concentration, b) luminescence and c) luminescence viable cell-1 following introduction of P. fluorescens into small pores (neck diameter < 6 µm) (o, •) or intermediate-sized pores (neck diameter 6-30 µm) (❏, ■) and introduction of sterile water (o, ❏) or C. steinii (•, ■) into large pores (neck diameter 30-60 µm). Error bars are minimum significant difference values calculated from two way analysis of variance for p ≤ 0.05. Bars are omitted where treatments were not significantly different (p > 0.05). Reprinted with permission from Wright D, Killham K, Glover LA et al. Appl Environ Microbiol 1995; 61:3537-3543.

5.5.6. Biosensors Use of living cells as biosensors offers several advantages over enzyme-based or other biosensors. Most importantly, analytical systems that require a sequence of biochemical reactions are greatly simplified by using cells within which all the reactions are conveniently packaged, and carried out efficiently in an optimal environment. In addition, production of bacterial-based biosensors is easily scaled-up, as only minor down-stream processing is needed after bacterial cultivation. Luminescence-based microbial biosensors for total toxicity have been developed and commercialized using naturally luminescent bacteria 49 and measure toxicity as quenching of luminescence following exposure of cells to a sample. Response times are short, ranging from few seconds50 to tens of minutes and operation is simple and does not require highly trained personnel or expensive instrumentation. Use of recombinant luminescent strains for general toxicity testing is now under evaluation for commercial products.51 One example is the assessment of inhibition of protein synthesis by strains of E. coli marked with prokaryotic or eukaryotic luciferase, using the strong bacteriophage λ leftward promoter.52 This provides several-hundred-fold induction in less than ten minutes and incubation prior to and after induction of protein synthesis enables distinction between effects on protein synthesis and direct metabolic inhibition. The test can be performed with lyophilized bacteria in less


than an hour and no bacterial cultivation is required, making it suitable for rapid and extremely sensitive detection of environmental samples.

5.6.1. Heavy Metal Sensors Regulatory elements from bacteria resistant to a heavy metal can provide a sensitive and selective receptor for heavy metal bioavailability53, 54 which will function in physiological concentration ranges. Genetic determinants for heavy metal resistance are usually found on plasmids and transposons of soil bacteria, which facilitate their analysis and manipulation by molecular genetic techniques. Many have been characterized, including mercury,55 arsenite,56 cadmium,57,58 zinc,59 cobalt,59 chromate,60 and copper.61 The first specific biosensor for bioavailable mercury was constructed utilizing firefly luciferase as a reporter gene.62 The regulatory unit of the mercury resistance operon from plasmid R100 was introduced into a plasmid vector pCSS810 21 to control the expression of firefly luciferase. Detection by this sensor strain, E. coli/pTOO11, gave a detection limit of approximately 1 fM HgCl2 (0.2 ppt Hg) and luminescence increased with increasing Hg (II) concentration. Selectivity was high, with interference only from cadmium at concentrations 108-fold higher than mercury. Sensitivity was sufficiently low for measurement of mercury in unpolluted natural waters, in which the concentration range is 2 - 60 ppt Hg.63 The detection limit of the routine method is approximately 200 ppt. A similar mercury sensor based on the usage of luciferase genes of Vibrio harveyi had a detection limit for mercury of 10 nM.64 Heavy metal sensors therefore operate at concentration ranges that are relevant to metal contamination of the environment, and that will be very useful in the future. An important feature is their amenability to lyophilisation, ensuring long shelf life (up to several years) and reagent-like usage.65, 66

5.6.2. Stress and Xenobiotics Sensors Bacteria in natural environments must protect themselves from a range of toxic insults, e.g., increased temperature, ultraviolet light, oxidative stress, redox-cycling xenobiotics and heavy metals. Protective mechanisms are best understood in E. coli in which exposure of aerobically growing cells to redox-cycling agents induces roughly 80 proteins.67 Stress response promoters have been utilized to generate stress biosensors. For example, plasmids have been constructed in which DNA damage-inducible promoters recA, uvrA, and alkA from E. coli have been fused to the V. fischeri luxCDABE operon.68 This allowed detection of a dose-dependent response to DNA-damaging agents, such as mitomycin and UV irradiation. A panel of such strains of lux biosensors may have use in monitoring chemical, physical, and genotoxic agents as well as in further characterizing the mechanisms of DNA repair. Individual strains from this panel will exhibit differential responses to specific stress conditions dependent on the regulatory circuit controlling the expression.69 A tod-luxCDABE fusion has been constructed and introduced into the chromosome of Pseudomonas putida F1.70 This strain was used to examine the induction of the tod operon when exposed to benzene, toluene, ethylbenzene and xylene (BTEX) compounds and aqueous solutions of JP-4 jet fuel constituents. Since this system contained the complete lux cassette (luxCDABE), bacterial bioluminescence in response to putative chemical inducers of the tod operon was measured on-line in whole cells without added aldehyde substrate. The sensor cells described above, together with specific heavy metal sensing strains, could be immobilized to an array of sensing elements and connected to a sensitive detector. This kind of biosensor might provide a more definitive understanding on the contents of complex environments through on-line monitoring of, for example, paper mill effluents and waste-water treatment plants.

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5.7. Conclusion All molecular marker systems enable the detection of tagged organisms through gene probing and viable cell enumeration. The major additional advantages offered by luminescence markers lie in the ability to detect light emission by marked organisms. This provides a degree of selective enumeration by colony counting, but is particularly valuable as a means of assessing real time, in situ metabolic activity of marked organisms, for instance in studies of spatial organization of organisms against background indigenous populations. Both prokaryotic and eukaryotic systems may be exploited and a range of vectors for each is available. Luminescence genes have also been used for assessment of general metabolic activity and as reporters of specific gene function. A range of relatively inexpensive methods is available for detection and luminescence-based markers have been used to study a variety of organisms and environments. The approach has particular advantages for monitoring of genetically engineered microorganisms released into the environment and, increasingly, in biosensor applications, and represents a powerful technique for fundamental studies of microbial ecology.

Acknowledgments MK acknowledges support from Maj and Tor Nessling Foundation. AJP acknowledges support to Spanish Ministry of Education, Junta de Andalucia and Fundación Ramón Areces. All authors are members of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Engebrecht J, Silverman M. Identification of genes and gene products necessary for bacterial bioluminescence. Proc Natl Acad Sci USA 1984; 81:4154-4158. 2. Eberhard A, Hastings JW. A postulated mechanism for the bioluminescent oxidation of reduced flavin mononucleotide. Biochem Biophys Res Comm 1972; 47:348-353. 3. Kurfurst M, Ghisla S, Hastings, JW. Characterization and postulated structure of the primary emitter in the bacterial luciferase reaction. Proc Natl Acad Sci USA 1984; 81:2990-2994. 4. Devine JH, Countryman C, Baldwin TO. Nucleotide sequence of the luxR and luxI genes and structure of the primary regulatory region of the lux regulon of Vibrio fischeri ATCC7744. Biochem 1988; 27:837-842. 5. Meighen EA. Molecular biology of bacterial bioluminescence. Microbiol Rev 1991; 55:123-142. 6. Soly RR, Mancini JA, Ferri, SR et al. A new lux gene in bioluminescent bacteria codes for a protein homologous to the bacterial luciferase subunits. Biochem Biophys Res Comm 1988; 155:351-358. 7. Baldwin TO, Devine JH, Heckel RC et al. The complete nucleotide sequence of the lux regulon of Vibrio fischeri and the luxABN region of Photobacterium leiognathi and the mechanism of control of bacterial bioluminescence. J Biolum Chemilum 1989; 4:326-341. 8. Swartzman E, Miyamoto C, Graham A et al. Delineation of the transcriptional boundaries of the lux operon of Vibrio harveyi demonstrates the presence of two new lux genes. J Biol Chem 1990; 265:3513-3517. 9. Meighen EA, Szittner RB. Multiple repetitive elements and organization of the lux operons of luminescent terrestrial bacteria. J Bacteriol 1992; 174:5371-5381. 10. Ulitzur S, Hastings JW. Evidence for tetradecanal as the natural aldehyde in bacterial bioluminescence. Proc Natl Acad Sci USA 1979; 76:265-267. 11. Riendeau D, Rodriguez A, Meighen, E. Resolution of the fatty acid reductase from Photobacterium phosphoreum into acyl protein synthetase and acyl-CoA reductase activities. J Biol Chem 1982; 257:6908-6915. 12. Rodriguez AL, Reindeau D, Meighen E. Fatty acid acylation of proteins in bioluminescent bacteria. Biochem 1983; 22:5604-5611.

Luminescence-Based Microbial Marker Systems and Their Application in Microbial Ecology 83 13. Byers DM, Meighen EA. Acyl-acyl carrier protein as a source of fatty acids for bacterial bioluminescence. Proc Natl Acad Sci USA 1985; 82:6085-6089. 14. Peabody DS, Andrews CL, Escudero, KW et al. A plasmid vector and quantitative techniques for the study of transcription termination in Escherichia coli using bacterial luciferase. Gene 1989; 75:289-296. 15. Engebrecht J, Simon M, Silverman M. Measuring gene expression with light. Science 1985; 227:1345-1347. 16. Lorenzo V, Herrero M, Jakubzik U et al. Mini-Tn5 transposon derivatives for insertion mutagenesis, promoter probing, and chromosomal insertion of cloned DNA in gram-negative eubacteria. J Bact 1990; 172:6568-6572. 17. Shaw J J, Settles LG, Kado CI. Transposon Tn4431 mutagenesis of Xanthomonas campestris pv. campestris: Characterization of a nonpathogenic mutant and cloning of a locus for pathogenicity. Mol Plant-microbe Interactions 1988; 1:39-45. 18. Wolk CP, Cai Y, Panoff, J-M. Use of a transposon with luciferase as a reporter to identify environmentally responsive genes in a cyanobacterium. Proc Natl Acad Sci USA 1991; 88:5355-5359. 19. Boivin R, Chalifour F-P, Dion P. Construction of a Tn5 derivative encoding bioluminescence and its introduction in Pseudomonas, Agrobacterium and Rhizobium. Mol Gen Genet 1988; 213:50-55. 20. Winson MK, Swift S, Hill PJ et al. Engineering the luxCDABE genes from Photorhabdus luminescens to provide a bioluminescent reporter for constitutive and promoter probe plasmids and mini-Tn5 constructs. FEMS Microbiol Lett 1998; 163:193-202. 21. Lampinen J, Koivisto L, Wahlsten M et al. Expression of luciferase genes from different origins in Bacillus subtilis. Mol Gen Genet 1992; 232:498-504. 22. Seliger HH, McElroy WD. Spectral emission and quantum yield of firefly bioluminescence. Arch Biochem Biophys 1960; 88:136-141. 23. Seliger HH, McElroy WD, White EH et al. Stereospecificity and firefly luminescence, a comparison of natural and synthetic luciferin. Proc Natl Acad Sci USA 1961; 47:1129-1134. 24. Wood KV, de Wet JR, Dewji N et al. Synthesis of active firefly luciferase by in vitro translation of RNA obtained from adult lanterns. Biochem Biophys Res Commun 1984; 124:592-596. 25. McElroy WD, DeLuca, M. Chemistry of firefly luminescence. In: Herring PJ ed. Bioluminescence in Action. Academic Press, London., 1978:109-127. 26. Wood KV, DeLuca M. Photographic detection of luminescence in Escherichia coli containing the gene for firefly luciferase. Anal Biochem 1987; 161:501-507. 27. Loimaranta V, Tenovuo J, Koivisto L et al. Generation of bioluminescent Streptococcus mutans and its usage in rapid analysis of antimicrobial compounds. Antimicrob Agents Chemother 1998; 42:1906-1910. 28. Kajiyama N, Nakano E. Enhancement of thermostability of firefly luciferase from Luciola lateralis by a single amino acid substitution. Biosci Biotechnol Biochem 1994; 58:1170-1171. 29. Denhardt DT, Colasanti J. A survey of vectors for regulating expression of cloned DNA in E. coli. In: Rodriguez RL, Denhardt DT eds. Vectors: a survey of molecular cloning vectors and their use. Butterworth Publishers, Stoneham, Mass. 1987:179-203. 30. Cebolla A, Vázquez ME, Palomares AJ. Expression vectors for the use of eukaryotic luciferases as bacterial markers with different colors of luminescence. Appl Environ Microbiol 1995; 61:660-668. 31. Vázquez ME, Cebolla A, Palomares, AJ. Controlled expression of click beetle luciferase using a bacterial operator-repressor system. FEMS Microbiol Lett 1994; 121:11-18. 32. Grant FA, Glover LA, Killham K et al. Luminescence-based viable cell enumeration of Erwinia carotovora in the soil. Soil Biol Biochem 1991; 23:1021-1024. 33. Wood KV, Lam YA, Seliger, HH et al. Complementary DNA coding click beetle luciferases can elicit bioluminescence of different colors. Science 1989; 244:700-702. 34. Shaw JJ, Dane F, Geiger D et al. Use of bioluminescence for detection of genetically engineered microorganisms released into the environment. Appl Environ Microbiol 1992; 58:267-273.

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35. Cebolla A, Ruiz F, Palomares AJ. Expression and quantification of firefly luciferase under control of Rhizobium meliloti symbiotic promoters. J Biolumin Chemilumin 1991; 6:177-184. 36. Cebolla A, Ruiz F, Palomares AJ. Stable tagging of Rhizobium meliloti with firefly luciferase gene for environmental monitoring. Appl Environ Microbiol 1993; 59, 2511-2519. 37. Coronado C, Vázquez ME, Cebolla A et al. Use of firefly luciferase gene for plasmid copy number determination. Plasmid 1994; 32:336-341. 38. Amin-Hanjani S, Meikle A, Glover LA et al. Plasmid and chromosomally encoded luminescence marker systems for detection of Pseudomonas fluorescens in soil. Mol Ecol 1993; 2:47-54. 39. Meikle A, Killham K, Prosser JI et al. Luminometric measurement of population activity of genetically modified Pseudomonas fluorescens in the soil. FEMS Microbiol Lett 1992; 99:217-220. 40. Duncan S, Glover LA, Killham K et al. Luminescence-based detection of activity of starved and viable but nonculturable bacteria. Appl Environ Microbiol 1994; 60:1308-1316. 41. Meikle A, Glover LA, Killham K et al. Potential luminescence as an indicator of activation of genetically modified Pseudomonas fluorescens in liquid culture and in soil. Soil Biol Biochem 1994; 26:747-755. 42. Langridge WHR, Fitzgerald KJ, Koncz C et al. Dual promoter of Agrobacterium tumefaciens mannopine synthase genes is regulated by plant growth hormones. Proc Natl Acad Sci USA 1989; 86:3219-3223. 43. O’Kane DJ, Lingle WL, Wampler JE et al. Visualization of bioluminescence as a marker of gene expression in Rhizobium infected soybean root nodules. Pl Mol Biol 1988; 10:387-399. 44. Herrero M, de Lorenzo V, Timmis K. Transposon vectors containing non antibiotic resistance selection markers for cloning and stable chromosomal insertion of cloned genes in gram-negative bacteria. J Bacteriol 1990; 172:6557-6567. 45. Palomares AJ, Vázquez ME, Cebolla A. Determination of Rhizobium nodule occupancy by luminescence. In: Hasting JW, Kricka LJ, Stanley PE eds. Bioluminescence and Chemiluminescence. Molecular Reporting with Photons. John Wiley and Sons, New York. 1997: 383-386. 46. Vázquez ME, Palomares AJ. Eukaryotic luciferase markers as monitoring methods for plasmid transfer analysis in soil environment In: Hasting JW, Kricka LJ, Stanley PE eds. Bioluminescence and Chemiluminescence. Molecular Reporting with Photons. John Wiley and Sons, New York. 1997:387-390. 47. Meikle A, Amin-Hanjani S, Glover LA et al. The effect of matric potential on survival and activity of a Pseudomonas fluorescens inoculum in soil. Soil Biol Biochem 1995; 27:881-892. 48. Wright D, Killham K, Glover LA et al. The role of pore size location in determining bacterial activity during protozoal predation. Appl Environ Microbiol 1995; 61:3537-3543. 49. Ribo JM, Kaiser KLE. Photobacterium phosphoreum, toxicity bioassay. Tox Assessm 1987; 2:305-323. 50. Lappalainen J, Juvonen R, Vaajasaari K et al. A new flash method for measuring the toxicity of solid and colored samples. 1998; Chemosphere:38,1069-1083. 51. Lampinen J, Virta M, Karp M. Use of controlled luciferase expression for monitoring of chemicals affecting protein synthesis. Appl Environ Microbiol 1995; 61:2981-2989. 52. Lampinen, J, Virta M, Karp, M. Comparison of gram-negative and grampositive bacterial strains cloned with different types of luciferase genes in bioluminescence cytotoxicity tests. Environ Toxicol Wat Qual 1995; 10:157-166. 53. O’Halloran TV. Transition metals in control of gene expression. Science 1993; 261:715-725. 54. Silver S, Walderhaug M. Gene regulation of plasmid- and chromosome-determined inorganic ion transport in bacteria. Microbiol Rev 1992; 56:195-228. 55. Summers AO. Organization, expression, and evolution of genes for mercury resistance. Ann Rev Microbiol 1986; 40:607-34. 56. Silver SK, Budd KM, Leahy WV et al. Inducible plasmid-determined resistance to arsenate, arsenite and antimony(III) in Escherichia coli and Staphylococcus aureus. J Bacteriol 1981; 146:983-996.

Luminescence-Based Microbial Marker Systems and Their Application in Microbial Ecology 85 57. Nucifora G, Chu L, Misra TK et al. Cadmium resistance from Staphylococcus aureus plasmid pI258 cadA gene results from cadmium-efflux ATPase. Proc Natl Acad Sci USA 1989; 86:3544-3548. 58. Tauriainen S, Karp M, Chang W et al. Luminescent bacterial sensor for cadmium and lead. Biosensors Bioelectronics 1998; 13:931-938. 59. Nies DH, Silver S. Plasmid-determined inducible efflux is responsible for resistance to cadmium, zinc, and cobalt in Alcaligenes eutrophus. J Bacteriol 1989; 171:896-900. 60. Peitzsch N, Eberz G, Nies DH. Alcaligenes eutrophus as a bacterial chromate sensor. Appl Environ Microbiol 1998; 64:453-458. 61. Cha J-S, Cooksey DA. Copper resistance in Pseudomonas syringae mediated by periplasmic and outer membrane proteins. Proc Natl Acad Sci USA 1991; 88:8915-8919. 62. Virta M, Lampinen J, Karp M. A luminescence-based mercury biosensor. Anal Chem 1995; 34:667-669. 63. Tescione L, Belfort G. Construction and evaluation of a metal ion biosensor. Biotechnol Bioeng 1993; 42:945-952. 64. Condee CW, Summers AO. A mer-lux transcriptional fusion for real-time examination of in vivo gene expression kinetics and promoter response to altered superhelicity. J Bacteriol 1992; 174:8094-8101. 65. Tauriainen S, Karp M, Chang W et al. A luminescence-based biosensor for arsenite and antimonite. Appl Environ Microbiol 1997; 63:4456-4461. 66. Ramanathan S, Ensor M, Daunert S. Bacterial biosensors for monitoring toxic metals. Trends Biotechnol 1997; 15:500-506. 67. Greenberg JT, Demple D. A global response induced in Escherichia coli by redox-cycling agents overlaps with that induced by peroxide stress. J Bacteriol 1989; 171:3933-3939. 68. Vollmer AC, Belkin S, Smulski DR et al. Detection of DNA damage by use of Escherichia coli carrying recA’::lux, uvrA’::gram, or alkA’::lux reporter plasmids. Appl Environ Microbiol 1997; 63:2566-2571. 69. Ben-Israel O, Ben-Israel H, Ulitzur S. Identification and quantification of toxic chemicals by use of Escherichia coli carrying lux genes fused to stress promoters. Appl Environ Microbiol 1998; 64:4346-4352. 70. Applegate, BM, Kehrmeyer SR, Sayler GS. A chromosomally based tod-luxCDABE wholecell reporter for benzene, toluene, ethylbenzene, and xylene (BTEX) sensing. Appl Environ Microbiol 1998; 64:2730-2735.

CHAPTER 6

The Use of the GUS Reporter System to Study Molecular Aspects of Interactions Between Bacteria and Plants M. Lambrecht, A. Vande Broek and J. Vanderleyden

6.1. Introduction

T

he most important criterion for an ideal marker/reporter system for rhizosphere bacteria is the absence of endogenous activity in the microbe of interest and in its natural environment. Other criteria include inexpensive use of substrates and/or material necessary to detect the marker, the availability of simple though sensitive, nondestructive, and histochemical assays to detect the strain or gene activity in planta, and unaffected fitness of the host after introduction of the marker.1 Although lacZ,2 encoding β-galactosidase, and phoA,3 encoding alkaline phosphatase, are well suited as gene fusion markers and have been used in plant-interacting bacteria, a high background in rhizosphere bacteria and higher plants often restricts their use in in planta studies. Luciferase activity can be detected using sensitive and nondestructive assays but needs costly materials and relies on the presence of oxygen and reduced cofactors in the host cell.4 The major advantage of the bioluminescent system is the total absence of activity in the plant-microbe background (see Chapter 5).5 The InaZ reporter (bacterial ice nucleation protein) does not need substrates nor sample processing for detection.6 Its major advantage is a logarithmic relationship between ice nucleation activity and InaZ protein concentration. However, it remains difficult to relate reporter activity to actual promoter strength or gene transcriptional activity. The xylE gene product, catechol-2,3-dioxygenase,7 is detected by means of a soluble compound and therefore not suitable to detect bacteria attached to plant tissue.8 Recently, the gfp gene, encoding green fluorescent protein, came into use as a reporter gene in diverse biological systems (see Chapter 7).9 The E. coli gusA gene (previously also named uidA), encoding β-glucuronidase, was first used as a reporter gene in plants.10,11 It is currently being used in a wide variety of biological systems, including plant-interacting bacteria. GUS activity is not present in most higher plants12 and in bacteria of agricultural importance such as Meso-, Sino-, Azo-, Brady Rhizobium, Agrobacterium, Azospirillum and Pseudomonas.13 In this chapter, the GUS reporter system and its use to study bacterium-plant interactions are reviewed. Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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6.2. The Escherichia coli gusA Gene Originally, β-glucuronidase was biochemically characterized in the bacterium Escherichia coli14 and later the gusA gene was isolated from E. coli strain K12.15 GUS activity enables E. coli in its natural environment, the gut, to decouple glucuronic acid from various substrates13 and to use glucuronic acid for further metabolization. The compounds coupled to glucuronic acid, the aglycones, can be very diverse. They are coupled to glucuronic acid in the liver to make them more water-soluble and are taken up again after decoupling by E. coli. This circulation is called the enterohepathic cycle, an important physiological process in the human body. In E. coli, gusA is part of an operon, together with two other genes, gusB and gusC, encoding a glucuronide-specific permease and a membrane-associated protein of unknown function, respectively (Fig. 6.1). The E. coli gusA gene is 1809 bp long and the β-glucuronidase has a predicted monomer molecular weight of 68,300, in agreement with the experimentally determined molecular weight of 73,000.15 The enzyme is probably active as a tetramer in vivo.11 Genetic analysis of the gusA locus has revealed three distinct modes of transcriptional control in E. coli. Firstly, the GusR repressor binds to an operator sequence upstream of gusA and is relieved from binding in the presence of glucuronide substrates. A second repressor, UxuR, downregulates both the gusA gene and the uxuAB operon,16,17 required for further metabolization of the glucuronic acid. A third level of control is exerted by catabolite repression.15 The β-glucuronidase of E. coli specifically hydrolyzes β -linked D -glucuronides to D-glucuronic acid and aglycone. The enzyme has a broad specificity range for β-conjugated glucuronides but not for other glycosides.18 β-glucuronidase was found to be very stable and to be more active in the presence of thiol-reducing agents such as β-mercaptoethanol and dithiothreitol (DTT).11 The optimum pH for GUS activity is between 5.0 and 6.8, but the enzyme retains about 50% activity at pH 4.3 or at pH 8.4. GUS is partially resistant to

Fig. 6.1. The gus operon in Escherichia coli.

The GUS Reporter System

89

Table 6.1. Characteristics of Escherichia coli β-glucuronidase (see text for ref.) Characteristics

Description

Substrates

β-linked D-glucuronides are hydrolized to D-glucuronic acid and aglycone No cleavage of glucuronides in internal positions within polymers (plant cell wall) Theoretical 68,000 ; experimental 72,000 5.0–7.0 (activity is 50% at pH 4.3 and 8.5) None Half-life of two hours at 55˚C and of 15 minutes at 60˚C Enhanced activity in presence of thiol reducing agents (DTT)

Exo-hydrolase Molecular weight pH range Cofactors Thermal stability Special requirements

thermal inactivation with a half-life of two hours at 55˚C and about 15 minutes at 60˚C. GUS is inhibited by some heavy divalent metal ions such as Cu2+ and Zn2+.11 Physical and enzymatic characteristics of GUS are summarized in Table 6.1. The enzyme can tolerate large amino-terminal additions for translational fusions. It is not processed at the amino-terminus in E. coli and is found exclusively in the cytoplasm. It is not subject to any post-transcriptional modification.

6.3. The GUS Reporter system Going back to the landmark publication of Jefferson et al,15 we can only now put in perspective to what extent the use of the Escherichia coli gusA gene as a reporter shifted the balance from reporter systems suited only for specific studies to a broad purpose, easy-touse, and precise reporter of gene expression. The use of fusions between a gene of interest and a reporter gene with an easily detectable phenotype, such as gusA, offers several advantages for the study of gene expression. Two types of gene fusions can be made. Transcriptional fusions are generated using sequences containing the coding sequence of GUS including its translational initiation signals. Expression of a transcriptional fusion results in the production of wild type β-glucuronidase. In a translational fusion, the translational initiation signal and the amino terminal coding sequences of a gene are linked in frame 5' to the coding sequence of gusA. Expression of a translational fusion results in the production of a chimeric protein. Fusion proteins are formed only when the reporter gene is inserted in the proper reading frame. Fusions can be generated in vivo by promoter probe transposons or in vitro by using promoter probe vectors. GUS can tolerate large amino-terminal fusions without loss of enzyme activity.10,15 While the data generated with GUS fusions to 5' control sequences of higher plant genes give only partial information of the transcriptional control of the gene of interest,19 this problem is not significant in prokaryotes. Besides E. coli, other bacteria, including soil bacteria, appear to have GUS activity.13 Plants grown in nonsterile soil occasionally exhibit GUS activity resulting from plant-associated bacteria. The proportion of soil bacteria expressing either LAC (β-galactosidase) activity or GUS activity has been estimated to be the same.20 It is not known to which degree the endogenous GUS activity of soil bacteria is inducible by the presence of glucuronides. Wilson et al13 found that some of the GUS activity in soil bacteria is inducible and may thus be responsible for some of the background activity when samples are incubated for a long period of time in the presence of GUS substrates.

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β-glucuronidase has become a widely used reporter system to measure expression of diverse genes by means of gene fusions (see 6.4). The composition of a typical extraction buffer used in quantitative GUS assays is 50 mM Na 2HPO4 (pH 6.0), 10 mM DTT or β-mercaptoethanol, 1 mM Na 2EDTA (to prevent inhibition by divalent metal ions), 0.1% Sodium Lauryl Sarcosine, 0.1% Triton X-10011 and substrate. Various fluorogenic and colorigenic substrates have been synthesized allowing quantification of the gene expression level in relatively simple and direct assays. para-Nitro-phenyl-β-D-glucuronide is a standard colorigenic substrate used for detection of β-glucuronidase activity in vitro.21 The liberated 4-nitrophenol can be measured spectrophotometrically at around 410 nm. The most frequently used fluorogenic substrate is 4-methylumbelliferyl-β-D-glucuronide (4-MUG). 4-methylumbelliferone formed by β-glucuronidase action has an excitation wavelength at 363 nm and an emission wavelength at 447 nm. Background fluorescence from the substrate is negligible.21 The product gives maximum fluorescence in the protonated phenoxide form and weak fluorescence in the phenol form. The equilibrium between these two forms is shifted towards the phenoxide form at pH values higher than its pKa value (for MUG, pKa 8.2). This results in rather low fluorescence at physiological pH values. Alternative GUS fluorochromes are available. For example, resorufin-β-D-glucuronide is hydrolyzed by GUS to resorufin. Resorufin, a phenoxazine derivative, has a high extinction coefficient and quantum efficiency, and excitation and emission maxima at wavelengths where plant tissue neither absorbs nor emits light. 4-Trifluoromethylumbelliferyl-β-D-glucuronide is a very sensitive substrate with emission maxima around 500 nm at which almost no plant tissue compounds emit energy. 3-Substituted umbelliferones such as 3-cyanoumbelliferyl-β-D-glucuronide have a reduced pKa in comparison to MUG. When hydrolyzed by GUS, these substrates thus have a stronger fluorescence at neutral pH, which can be an advantage for some assays. Fluorescein glucuronides are hydrolyzed to fluorescein by the GUS enzyme. Fluorescein is the most widely used fluorochrome in biology. It fluoresces very well in living cells although the main disadvantage of fluorescein glucuronides is the intr insic fluorescence of the substrates. Although the sensitivity of fluorogenic GUS assays is higher than the spectrophotometric assays, expensive fluorescence instrumentation is required when using these assays. The main advantage of fluorescence assays is that they can provide two or three orders of magnitude greater sensitivity than spectrophometric assays. Fluorescence assays do require slightly higher skills than spectrophotometric methods due to the attention which has to be given to control experiments for calibration of fluorometers, nonenzymatic turnover of fluorescent substrates in certain conditions and quenching of the fluorescence. The main benefit when using the GUS reporter system in microbial rhizosphere ecology is the ability to localize GUS marked bacteria on plants using histochemical GUS substrates. The colorless histochemical substrate X-GlcA (5-bromo-4-chloro-3-indolyl-β-D-glucuronide) is hydrolyzed by GUS to a blue indigo dye precipitate.21 After enzymatic reaction, the released indoxyl derivative must undergo an oxidative dimerization to form the insoluble and highly colored indigo dye.11 The dimerization is enhanced in the presence of atmospheric oxygen and can be stimulated by using an oxidative catalyst such as a ferricyanide/ferrocyanide mixture (K3Fe(CN)6 and K4Fe(CN)6 respectively). Naphtol AS-BI β-D-glucuronide is cleaved by GUS to liberate free naphtol AS-BI which must then be coupled to a diazo dye to give a precipitating azo dye. The assay has the capacity to generate different colors depending on the coupling dye used.22 Besides X-GlcA, other histochemical substrates have recently been synthesized20 which render products with various other colors (Table 6.2). In Table 6.3, an overview of constructs, containing the E. coli gusA gene, is given. The table is not exhaustive and only a selection of constructs with potential use in plant-microbe interaction studies, is mentioned.


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Table 6.2. Currently available histochemical GUS substrates15,20 Name

Chemical Name

Color of product

Company

X-GlcA

5-bromo-4-chloro-3indolyl-β-D-glucuronide 6-bromo-2-hydroxy-3naphthoyl-O-anisidineβ-D-glucuronide 5-bromo-6-chloro-3indolyl-β-D-glucuronide 6-chloro-3-indolyl-βD-glucuronide Indoxyl-β-D-glucuronide 6-chloro-3-indlyl-β-Dglucuronide

Blue

Biosynth AG, Research Organics Inc. CalBioChem Sigma

Naphtol ASBIglucuronide Magenta-glcA Salmon-glcA Indoxyl-glcA Red-Gluc

Depends on choice of coupling dye Magenta

Biosynth AG

Salmon pink

Biosynth AG

Indigo blue Red

Biosynth AG Research Organics Inc.

6.3.1. The Tn5-gusA Promoter probe transposons of Sharma and Signer23 Sharma and Signer23 constructed Tn5-gusA promoter probe transposons to monitor expression of Rhizobium meliloti symbiotic genes within alfalfa nodules. For these transposons, a Tn5 derivative,24 in which 25 base pairs of IS50L are retained, was used. Tn5gusA1 is a transcriptional operon fusion transposon. The gusA gene is preceded by translation stop codons in all open reading frames and a Shine-Dalgarno sequence to allow initiation of gusA translation. Tn5-gusA2 is similar although one of the stop codons and the Shine-Dalgarno sequence have been omitted, making it ideal for the construction of translational gene fusions.

6.3.2. GUS Constructs of Wilson et al25 Wilson et al25 constructed vectors useful for manipulation of gusA and a whole series of minitransposons containing gusA as a reporter gene. The vectors allow construction of translational fusions to gusA in all three open reading frames. In the miniTn5, constructed by de Lorenzo et al26 and Herrero et al,27 transposase activity inherent to Tn5 has been deleted allowing a stable insertion. Transposase activity has, thus, to be provided in trans for transposition. Wilson et al 25 used the unique restriction sites of mini-Tn5 to insert gusA and some other genes. The construct mTn5SSgusA20 gives high constitutive expression of gusA in Rhizobium. mTn5SSgusA10 contains the lacIq gene and gusA under the control of the tac promoter and, therefore, expression is repressed until an inducer of the lac operon, such as IPTG, is added. This allows regulation of gusA gene expression after introduction in the host strain. Postponing high GUS expression until the time of the GUS assay reduces the stress imposed on the bacterium during the normal life-cycle. mTn5SSgusA40 is a promoter probe transposon useful to screen for promoters, e.g., those responding to specific environmental signals.

6.3.3. Other GUS Constructs Five gusA cassettes, uidA1, uidA2, uidA2-cat, uidA2-aadA and uidA2-aph, suitable for constructing transcriptional fusions,were described by Metcalf and Wanner.28 Three of them contain additional antibiotic resistance genes (see Table 6.3). uidA1, uidA2 and uidA2-aph cause nonpolar mutations after double homologous recombination into the host genome.


92

Table 6.3. Overview of GUS constructs used in studies of plant-bacteria interactions Name

Description

Properties

Ref.

Tn5-gusA1

Promoter probe transposon

23

Tn5-gusA2

Promoter probe transposon gusA-expressing transposon gusA-expressing transposon

Construction of transcriptional fusion; presence of Shine-Dalgarno consensus sequence and stop codons in all open reading frames Construction of translational fusions; no Shine-Dalgarno sequence gusA under the control of the nptII promoter Same as mTn5SsgusA20; restriction site of rare cutter for chromosomal mapping of Tn5 insertion location lacIq, encoding LAC repressor, is present and gusA gene under control of tac promoter; expression of gusA dependent on addition of IPTG Same as mTn5SSgusA10 but no lacIq present

mTn5SsgusA20 Tn5SSgusA21 mTn5SsgusA10

gusA-expressing transposon

mTn5SsgusA11

gusA-expressing transposon gusA-expressing transposon

mTn5SsgusA30 mTn5SsgusA31

gusA-expressing transposon

mTn5SsgusA40

Promoter probe transposon GUS expression vector

pRAJ289

pRAJ294 pKW117 pKW118 pKW119 pRG960

pRG970

pFAJ31 pWM2 (uidA1)

GUS expression vector Translation fusion vectors Broad host range and promoter selection cosmid

Broad host range and promoter selection cosmid GUS fusion vector contains gusA cassette for generating transcriptional fusions

gusA under the control of UAS and σ54-dependent sequence of Rhizobium etli CFN42 nifH gene promoter gusA under the control of σ54-dependent sequence of Bradyrhizobium sp. Parasponia strain Rp501 nifH gene promoter Promoterless gusA can be transcribed from adjacent promoters in the genome. gusABC and unknown ORF under the control of the tac promoter on a multicopy expression vector carrying lac repressor gene Same as pRAJ289 but only gusA present

23 25 25

25

25 25

25

25 25

25

Construction of translational fusion to gusA in all 3 open reading frames

25

Replication in wide variety of gramnegative bacteria ; compatible with IncP cloning vectors; construction of transcriptional and translational fusions to a promoterless gusA ; cos site Same as pRG960 ; isolation of promoter due to presence of one multiple cloning site upstream of both gusA and lacZ pLAFR3 derivative with promoterless gusA gene Non-polar mutation after allele replacement in the host genome ; no antibiotic resistance

29

29

31 28


93

Name

Description

Properties

pWM3 (uidA2)

contains gusA cassette for generating transcriptional fusions contains gusA cassette for generating transcriptional fusions contains gusA cassette for generating transcriptional fusions contains gusA cassette for generating transcriptional fusions Promoter-probe transposon Promoter-probe transposon

Same as pWM2 but different flanking multiple cloning site

28

Same as pWM3 but cat gene present in cassette; polar mutation after introduction in the host genome

28

Same as pWM3 but SpR and SmR genes present in cassette; polar mutation after introduction in the host genome

28

Same as pWM3 but AmpR gene present in cassette; nonpolar mutation after introduction in the host genome

28

pWM4 (uidA2-cat)

pWM5 (uidA2-aadA)

pWM6 (uidA2-aph) mTn5gfp-pgusA mTn5gusA-pgfp11

mTn5gusA-pgfp12 mTn5gusA-pgfp21 mTn5gusA-pgfp22

Promoter probe transposon Promoter probe transposon Promoter probe transposon

Ref.

Promoterless gfp followed by a 34 constitutively expressed gusA gene. Promoterless gusA preceded by consensus 34 Shine-Dalgarno sequence and translational stop codons in three ORF’s. Expression of uv-gfp controlled by nptII promoter Same as mTn5gusA-gfp11. Expression of 34 uv-gfp controlled by tandem nptII promoter Same as mTn5gusA-gfp11. Two copies of 34 gfp are each controlled by a nptII promoter Same as mTn5gusA-gfp11. Two copies of gfp 34 are each controlled by a tandem nptII promoter.

Several researchers constructed reporter vectors based on gusA and lacZ, facilitating the analysis of bidirectional promoter regions.29,30 All of these constructs can be used to monitor free-living populations of bacteria in soil and to monitor the colonization of plant roots by bacteria. Additionally, a number of constructs have been specifically developed to monitor the colonization of plant roots by Azospirillum31,32 and Azoarcus.33 Recently, a series of promoter-probe transposons, mTn5gusA-pgfp, containing a promoterless gusA gene and one or two constitutively expressed gfp genes has been constructed and found to be useful for gene expression studies in Rhizobium, Azospirillum and Pseudomonas.34

6.4 Application of the GUS Reporter System in Studies of Plant-Bacterium Interactions This section deals with studies of rhizosphere bacteria, i.e., Rhizobium, Azospirillum, Azoarcus, Xanthomonas and Pseudomonas, which have used GUS as a reporter. In these studies, GUS has been used to mark the soil bacterium of interest or to analyze the expression of a specific gene in planta. Only a few representative cases for the different genera are discussed.

6.4.1. Rhizobium-legume Symbiosis Sharma and Signer23 have used Tn5-gusA1 and Tn5-gusA2 to study the expression of Rhizobium meliloti symbiotic genes during nodule organogenesis of alfalfa. Two patterns of

94


spatial and temporal expression of the symbiotic genes were observed. The nod genes, involved in nodule formation, are expressed in an early phase (from 36 hours after inoculation) on the root surface and nodule cortex and later (from 10 days after inoculation) especially at the nodule meristem. In contrast, the nif (encoding nitrogenase), fix (required for nitrogen fixation) and syrM (a regulator of other symbiotic genes) are expressed 10 days after inoculation in the active symbiotic zone of the alfalfa nodules spreading off to the rest of the central region of the nodule. For histochemical assay of β-glucuronidase, nodules were embedded, fixed and stained with 20 µg/ml 1-GlcA. The transposons described by Wilson et al 25 were designed to monitor population changes in the soil and the rhizosphere and to determine nodule occupancy in rhizobial competition studies. All assay parameters to study rhizobial infection and nodule occupancy were optimized. The ideal concentration of the histochemical substrate X-Glc was 100 µg/ml. Elevating the substrate concentrations to 250 or 500 µg/ml elicited intrinsic and inducible GUS activity from other soil bacteria. Rhizobial cells, marked with mTn5SSgusA10, mTn5SSgusA11, mTn5SSgusA20 or mTn5SSgusA21, could be visualized especially on young nodules. Rhizobia marked with mTn5SSgusA30 or mTn5SSgusA31, containing gusA expressed from a nifH promoter, could be visualized in older nodules with maximal expression in the central, nitrogen-fixing zone. In Figure 6.2, an X-GlcA stained section of an alfalfa root nodule, infected with R. meliloti containing a nifH-gusA translational fusion, is shown.35 Streit et al36 used a GUS+ strain of R. leguminosarum bv. phaseoli strain KIM5s to test the nodulation competitiveness of 17 Rhizobium leguminosarum bv. phaseoli and 3 R. tropici strains. They found that strains which could use simple aromatic compounds were generally more competitive. Kalinowski and Long37 used fusions between deletion derivatives of a selected nod gene and gusA to monitor expression levels in free-living conditions and in planta.

6.4.2. Plant Growth Promoting Bacteria (PGPR) of the Genus Azospirillum Bacteria living in the rhizosphere and favorably affecting plant growth are denominated Plant Growth Promoting Rhizobacteria (PGPR). PGPR of the genus Azospirillum were initially studied because of their association with Gramineae and because of their ability to fix atmospheric nitrogen.38 Field trials, however, indicated that bacterial nitrogen is not responsible for the enhanced plant growth observed upon Azospirillum inoculation.39 The observation that inoculation of plant roots with Azospirillum results in a clear plant root proliferation intensified the study of bacterial phytohormone biosynthesis. For the azospirilla to exert their beneficial effect, it is well recognized that rhizosphere colonization is a prerequisite for enhancing plant growth. Aspects of bacterial rhizosphere colonization include chemotactic motility, adherence to the plant root surface and utilization of rhizosphere nutrients. The study of an associative plant-bacterium interaction is hampered by the lack of an easily detectable plant phenotype following inoculation. To visualize the Azospirillum-plant root association, A. brasilense strains carrying a constitutive gusA fusion (pFAJ31.13) were constructed.31 pFAJ31.13 was constructed by inserting an A. brasilense gene library upstream of a promoterless gusA gene and selecting a fusion for high β-glucuronidase activity under different physiological conditions. This fusion was then considered useful to localize Azospirillum on wheat roots in colonization assays. During the first days of the association, the bacterial cells were mainly found in the root hair zones (see Fig. 6.3) and at the sites of lateral root emergence. Microscopic analysis of sections through colonized zones occasion-


95 Fig. 6.2. Section of an X-GlucA stained alfalfa nodule, infected with R. meliloti (nifH-gusA)

ally revealed the presence of azospirilla in the epidermis and cortex layers. No bacterial penetration through the endodermis or vascular tissue was observed. In a second study, the colonization of wheat roots by a selection of A. brasilense mutants impaired in motility, chemotaxis and plant root attachment, carrying pFAJ31.13, was evaluated. Two nonmotile and a generally nonchemotactic mutant were found to be impaired in the primary colonization of the wheat roots. It can be hypothesized, that key compounds are specifically secreted by the root hair zone of the wheat roots to which Azospirillum is attracted.40,41 The expression of a translational A. brasilense nifH-gusA fusion was studied in free-living conditions and during wheat root association.31 Free-living nifH expression was shown to occur only under nitrogen-limiting microaerobic conditions, suggesting the presence of nitrogen and oxygen-dependent control mechanisms for nif gene expression in Azospirillum. Analysis of nifH gene expression and nitrogenase activity during the Azospirillum-wheat root association indicated that both oxygen and the availability of carbon sources are limiting factors for associative nitrogenfixation.

96


Fig. 6.3. Primary colonization of the wheat hair root zone by Azospirillum brasilense Sp245 (pFAJ31.13).

6.4.3. Root Colonization and Systemic Spreading of Azoarcus sp. Strain BH72 Azoarcus sp.42 is an endorhizospheric isolate of Kallar grass. Kallar grass is an undomesticated C4 plant with high tolerance to soil salinity, alkalinity and waterstress. Similarly as has been described for Azospirillum, colonization of Kallar grass by Azoarcus has been studied using a constitutively expressed gusA fusion. For this, mutants of Azoarcus were generated by transposon mutagenesis with Tn5-gusA1 (Table 6.1) and selected for constitutive GUS expression.33 Microscopic examination of Kallar grass treated with GUS-marked wild-type Azoarcus revealed bacterial colonization inside and between cortex cells of the roots. In particular, identity of the inoculant Azoarcus cells inside cells of roots was established by histochemical GUS staining.33

6.4.4. Plant-Pathogenic Bacteria GUS was used to study the expression of the sod gene encoding superoxide dismutase from Xanthomonas campestris pv. campestris both ex planta and in planta.43 The formation of active oxygen species (AOS) in plants is believed to be important in host defense mechanisms. Superoxide dismutase (SOD), among other microbial enzymes, protects the pathogenic bacterium from the O2- produced by the plant by reduction to H2O2 and O2. Expression of a X. campestris sod-gusA transcriptional fusion was monitored in bacterial strains released onto plants. A clear induction of the sod-gusA fusion in in planta experiments could be observed. The second burst of plant AOS production, typical for incompatible interactions, is stimulated only by bacteria that cause a hypersensitive response. However, no difference in GUS activity was measured following a compatible or an incompatible interaction of X. campestris on turnip and pepper, respectively. The gene was not induced by conditions mimicking the first AOS burst, occurring both in compatible and incompatible interactions. These two observations suggest that the induction of the X. campestris SOD enzyme under in planta conditions is not a direct response to extracellular O2- or H2O2.43 Xiao et al 44 performed site-directed Tn5-gusA1 mutagenesis on the hrp genes of Pseudomonas syringae pv. syringae. The hrp gene cluster functions in elicitation of the hypersensitive response when a nonhost plant species (or a resistant variety of a susceptible plant species) is inoculated with P. syringae and is also required during pathogenesis in a susceptible host. On the basis of the orientation of Tn5-gusA1 insertions and their relative GUS activities in planta, the transcriptional organization of the P. syringae pv. syringae hrp


97

gene cluster could be deduced. Since levels of expression were similar in minimal medium and during the early stages of interaction with tobacco cells, it was suggested that nutritional conditions, rather than a particular plant factor, affect expression of the hrp genes in P. syringae pv. syringae.

6.5 Conclusion The GUS system is a precise and robust reporter for gene expression, allowing transcriptional and translational fusions between the gene of interest and the gusA gene. The main advantages of GUS are the absence of endogenous activity in plants, the existence of a large number of commercially available substrates and the number of constructs facilitating the use of the E. coli gusA gene. GUS has been used both as a reporter and marker in studies with rhizosphere bacteria, e.g., the Rhizobium-legume symbiosis, PGPR, endophytic and phytopathogenic bacteria. Histochemical assays allow the assessment of the localization of GUS marked rhizosphere bacteria on plant roots. Quantitative assays exist to count the number of GUS marked bacteria in the rhizosphere. GUS-marked recombinant Azospirillum strains have been used in field release experiments in Italy in 1994 and 1995. The gusA gene, under the control of the nptII promoter, was inserted in the genome of Azospirillum brasilense (wild-type and strains altered in auxin biosynthesis). The engineered strains were used for inoculation of sweet sorghum by Agronomica (Italy) in collaboration with the University of Padua, in the framework of an EU project. The GUS marker allowed monitoring of survival and spread of the recombinant bacteria in soil samples with a detection limit of 102 CFUs of GUS-marked strains per gram soil.

Acknowledgments M.L. and A.V.B. are recipients of a predoctoral and postdoctoral fellowship of the Fund for Scientific Research, Flanders, respectively. J. Vanderleyden is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Wilson KJ. Molecular techniques for the study of rhizobial ecology in the field. Soil Biol Biochem 1995; 27:501-514. 2. Silhavy TJ, Beckwith JR. Uses of lac fusions for the study of biological problems. Microbiol Rev 1985; 49:398-418. 3. Reuber TL, Long, SL, Walker GC. Regulation of Rhizobium meliloti exo genes in free-living cells and in planta examined using TnphoA fusions. J Bacteriol 1991; 173:426-434. 4. de Weger L, Dekkers LC, van der Bij AJ et al. Use of bioluminescence markers to detect Pseudomonas spp. in the rhizosphere. Appl Environ Microbiol 1991; 57:3641-3644. 5. Cebolla A, Ruiz-Berrasuero F, Palomares AJ. Stable tagging of Rhizobium meliloti with the firefly luciferase gene for environmental monitoring. Appl Environ Microbiol 1993; 59:25112519. 6. Lindgren PB, Frederic R, Govindarajan AG. An ice nucleation gene reporter gene system identification of inducible pathogenicity genes in Pseudomonas syringae pv. phaseolicola. EMBO J 1989; 8:1291-1301. 7. Winstanley C, Morgan JA, Pickup R et al. Use of a xylE marker gene to monitor survival of recombinant Pseudomonas populations in lake water by culture on nonselective media. Appl Environ Microbiol 1991; 57:1905-1913. 8. Buell CR, Anderson AJ. Expression of the aggA locus of Pseudomonas putida in vitro and in planta as detected by the reporter gene xylE. Mol Plant-Microbe Int 1993; 6(3):331-340.

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9. Chalfie M, Tu Y, Euskirchen G et al. Green fluorescent protein as a marker for gene expression. Science 1994; 263:802-805. 10. Jefferson RA, Kavanagh TA, Bevan MW. GUS fusions : β-glucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO J 1987; 6(13):3901-3907. 11. Jefferson RA. Assaying chimeric genes in plants : The GUS gene fusion system. Plant Molecular Biology Reporter 1987; 5:387-405. 12. Hu CH, Chee PP, Chesney RH et al. Intrinsic GUS-like activities in seed plants. Plant Cell Reports 1990; 9:1-5. 13. Wilson K, Hughes SG, Jefferson RA. The Escherichia coli gus operon : introduction and expression of the gus operon in E. coli and the occurrence and use of GUS in other bacteria. In : Gallagher SR, ed. GUS protocols. Using the GUS gene as a reporter of gene expression. San Diego : Academic Press, Inc. 1992:7-22. 14. Stoeber F. Etudes des propriétés et de la biosynthèse de la glucuronidase et de la glucuronide-permease chez Escherichia coli. Thèse de Docteur des Sciences 1961; Université de Paris, France. 15. Jefferson RA, Burgess SM, Hirsh D. β-glucuronidase from Escherichia coli as a gene-fusion marker. Proc Natl Acad Sci USA 1986; 83:8447-8451. 16. Novel M, Novel G. Regulation of β-glucuronidase synthesis in Escherichia coli K-12 : constitutive mutants specifically derepressed for uidA expression. J Bacteriol 1976; 127:406-417. 17. Novel M, Novel G. Regulation of β-glucuronidase synthesis in Escherichia coli K-12 : pleiotropic constitutive mutations affecting uxu and uidA expression. J Bacteriol 1976; 127:418-432. 18. Tomasic J, Keglevic D. The kinetics of hydrolysis of synthetic glucuronic esters and glucuronic ethers by bovine liver and Escherichia coli β-glucuronidase. Biochem J 1973; 133:789. 19. Taylor CB. Promoter fusion analysis: An insufficient measure of gene expression. The Plant Cell 1997; 9:273-275. 20. Wilson KJ. gusA as a reporter gene to track microbes. In: Akkermans ADL, van Elsas JD, de Bruijn FJ, eds. Molecular Microbial Ecology Manual. Dordrecht : The Netherlands, 1995. 21. Naleway JJ. Histochemical, spectrophotometric, and fluorometric GUS substrates. In: Gallagher, SR, ed. GUS protocols. Using the GUS Gene as a Reporter of Gene Expression. San Diego ; Academic Press, Inc. 1992: 61-76. 22. Fishman W, Nakajima Y, Anstiss C et al. Napthol AS-BI β-D-glucuronidic acid: Its synthesis and suitability as a substrate for β -glucuronidase. J Histochem Cytochem 1964; 12:298-305. 23. Sharma SB, Signer ER. Temporal and spatial regulation of the symbiotic genes of Rhizobium meliloti in planta revealed by transposon Tn5-gusA. Genes Dev 1990; 4:344-356. 24. Simon R, Quandt J, Klipp W. New derivatives of transposon Tn5 suitable for mobilization of replicons, generation of operon fusions and indction of genes in gramnegative bacteria. Gene 1989; 80:161-169. 25. Wilson KJ, Sessitsch A, Corbo J et al. β-glucuronidase (GUS) transposons for ecological and genetic studies of rhizobia and other gramnegative bacteria. Microbiology 1995; 141:1691-1705. 26. de Lorenzo V, Herrero M, Jakubzik U et al. Mini-Tn5 transposon derivatives for insertion mutagenesis, promoter probing, and chromosomal insertion of cloned DNA in gramnegative eubacteria. J Bacteriol 1990; 172:6568-6572. 27. Herrero M, de Lorenzo V, Timmis KT. Transposon vectors containing non-antibiotic resistance selection markers for cloning and stable chromosome insertion of foreign genes in gramnegative bacteria. J Bacteriol 1990; 172:6557-6567. 28. Metcalf WW, Wanner BL. Construction of new β-glucuronidase cassettes for makiing transcriptional fusions and their use with new methods for allele replacement. Gene 1993; 129:17-25. 29. Van den Eede G, Deblaere R, Goethals K et al. Broad host range and promoter selection vectors for bacteria that interact with plants. Mol Plant-Microbe Interact 1992 ; 5(3):228-234. 30. Parry SK, Sharma SB, Terzaghi EA. Construction of a bidirectional promoter probe vector and its use in analysing nod gene expression in Rhizobium loti. Gene 1994; 150:105-109.


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31. Vande Broek A, Michiels J, Van Gool A et al. Spatial-temporal colonization patterns of Azospirillum brasilense on the wheat root surface and expression of the bacterial nifH gene during association. Mol Plant-Microbe Interact 1993; 2:261-266. 32. Christiansen-Weniger C, Vanderleyden J. Ammonium-excreting Azospirillum sp. become intracellularly established in maize (Zea mays) para-nodules. Biol Fertil Soils 1993; 17:1-8. 33. Hurek T, Reinhold-Hurek B, van Montagu M et al. Root colonization and systemic spreading of Azoarcus sp. strain BH72 in grasses. J Bacteriol 1994; 176:1913-1923. 34. Xi C, Lambrecht M, Vanderleyden J et al. Bi-functional gfp-and gusA containing mini-Tn5 transposon derivatives for combined gene expression and bacterial localization studies. J Microbiol Meth 1999; 35:85-92. 35. Vande Broek A, Michiels J, de Faria SM et al. Transcription of the Azospirillum brasilense nifH gene is positively regulated by NifA and NtrA and is negatively controlled by the cellular nitrogen status. Mol Gen Genet 1992; 232:279-283. 36. Streit W, Kosch K, Werner D. Nodulation competitiveness of Rhizobium leguminosarum bv. phaseoli and Rhizobium tropici strains measured by glucuronidase (gus) gene fusion. Biol Fertil Soils 1992; 14:140-144. 37. Kalinowski G, Long SR. Deletion analysis of the 5’ untranslated region of the Rhizobium meliloti nodF gene. Mol Plant-Microbe Interact 1996; 9:869-873. 38. Beijerinck MW. Über ein Spirillum, welche freien Stikstoff binden kann ? Zentralbl Bakteriol Parasitenkd Infektionskr Abt 1925; 63:353. 39. Okon Y, Vanderleyden J Root-associated Azospirillum species can stimulate plants. ASM News 1997; 63:364-370. 40. Vande Broek A, Vanderleyden J. The role of bacterial motility, chemotaxis, and attachment in bacteria-plant interactions. Mol Plant-Microbe Interact 1995; 8:800-810. 41. Vande Broek A, Lambrecht M, Vanderleyden J. Bacterial chemotactic motility is important for the initiation of wheat root colonization by Azospirillum brasilense. Microbiology 1998; 144:2599-2606. 42. Reinhold-Hurek B, Hurek T, Gillis M et al. Azoarcus gen. nov., nitrogen-fixing proteobacteria associated with roots of Kallar grass (Leptochloa fusca (L.) Kunth), and description of two species, Azoarcus indigens sp. nov., and Azoarcus communis sp. nov. Int J Syst Bacteriol 1993; 43:574-584. 43. Smith SG, Greer Wilson TJ, Maxwell Dow J et al. A gene for superoxide dismutase from Xanthomonas campestris pv. campestris and its expression during bacterial –plant interactions. Mol Plant-Microbe Interact 1996 ; 9:584-593. 44. Xiao Y, Lu Y, Heu S et al. Organization and environmental regulation of the Pseudomonas syringae pv. syringae 61 hrp cluster. J Bacteriol 1992; 174:1734-1741.

CHAPTER 7

Using Green Fluorescent Protein (GFP) as a Biomarker or Bioreporter for Bacteria J. R. Stoltzfus, J.K. Jansson and F.J. de Bruijn

7.1 Introduction

S

ince the cloning of the Green Fluorescent Protein (GFP) gene from the jellyfish Aequorea victoria,1 and its expression in other organisms,2 GFP has rapidly become an important biomarker/bioreporter in a wide variety of eukaryotic and prokaryotic organisms. The utility of GFP as a bioreporter/biomarker lies mainly in the simplicity of the biochemical reaction generating fluorescence. GFP expression and fluorescence does not depend on the addition of co-factors or additional substrates, and only requires oxygen briefly for the autooxidation of GFP. When GFP is excited with light of the proper wavelength it autofluoresces. Visualization of this fluorescence can be used to monitor gene expression, to localize proteins, to isolate novel genes, or to track tagged cells (For reviews see refs. 3-9). Because of these advantages, GFP is replacing some of the traditional biomarkers/bioreporters used in molecular microbial ecology, such as metabolic markers or luciferase markers which require substrate addition and cellular energy reserves for visualization (For reviews see refs. 6, 10, 11).

7.2 Properties of Wild-type GFP Wild-type GFP (wt GFP) is a 27 kDa monomer consisting of 278 amino acid residues.1 In Aequorea victoria, GFP serves to convert blue chemiluminescence, generated by the protein aequorin, into green light.12 This conversion is accomplished by means of an internal protein p-hydroxybenzylideneimidazolinone chromophore formed by the cyclization of three amino acid residues of GFP, Ser-Tyr-Gly, followed by oxidation of the Tyr residue. These essential three residues are located at positions 65-67 of the GFP protein.13 The formation and activation of the GFP chromophore does not require additional enzymes or cofactors, and therefore heterologous expression of GFP in numerous organisms has been shown to result in fluorescence.3 The wtGFP absorbs light with an excitation maximum of 395 nm, with a second excitation peak at 470 nm and it fluoresces with an emission maximum of 510 nm.14,15 The excitation/emission characteristics of GFP have important implications for its use as a biomarker/bioreporter. The 470 nm excitation peak allows observation of wtGFP fluorescence with a standard fluorescein isothiocyanate (FITC) excitation-emission filter set, or with the 488 nm line of an argon laser, equipped with appropriate filters. This allows the Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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detection of GFP using fluorescence-activated cell sorting (FACS), flow cytometry, epifluorescent microscopy and laser-scanning confocal microscopy (LSCM). The fluorescence emission of the wtGFP when excited with longer wavelength light is less intense than its emission when excited with shorter wavelength light. This renders detection of the low levels of fluorescence emitted from wtGFP expressed in some bacteria problematic. On the other hand, excitation of wtGFP with longer wavelength light causes less photobleaching than excitation with shorter wavelength light, making longer wavelength light a better choice for exciting GFP.2 One possible reason for the low fluorescence emission levels observed when wtGFP is expressed in selected bacteria is that much of the wtGFP produced is found in inclusion bodies in an insoluble non-fluorescent form.16 For example, populations of E. coli cells containing plasmids encoding wtGFP give rise to a non-fluorescent subpopulation, presumably due to formation of inclusion bodies.17,18 A similar effect has been observed in clonal populations of Pseudomonas putida tagged with wtGFP.19

7.3. Improving GFP Fluorescence GFP from the sea pansy Renilla reniformis contains the same chromophore as GFP from Aequorea victoria. However, the GFP from R. reniformis has a single excitation peak at 498 nm. This difference is presumably due to the difference in amino acid residues surrounding the chromophore in the mature protein.15 Neither the isolated chromophore, nor the denatured protein at neutral pH, are fluorescent.13 These data suggested that changing the protein environment (amino acid sequence) around the chromophore of GFP could change the excitation/emission characteristics of GFP. A number of methods to generate GFP mutants with altered excitation-emission spectra and enhanced emission intensities have been used to increase the utility of GFP as a biomarker/ bioreporter (See Table 7.1). For example, hydroxylamine treatment and error-prone PCR were used to generate mutant cDNAs encoding new versions of GFP.16 Colonies of E. coli harboring mutant cDNAs were screened for changes in emission color and brightness when excited with 375 vs 495 nm light. Five mutants were identified in this screen. Three of these mutants (H9, P9 and P11, Table 7.1) displayed significant alterations in excitation maxima, with little or no effect on emission spectra, and had an enhanced fluorescence level. None of these three mutants contained changes in the chromophore-forming residues. The other two mutants, P4 and W, both harbored substitutions at position 66, which forms part of the chromophore, and exhibited blue fluorescence. In addition, the excitation maxima of mutant W was increased (Table 7.1). Site-directed mutagenesis of the serine residue at position 65 yielded four mutants of GFP with single excitation peaks in the 470-490 nm range and an enhanced level of light emission.20 The S65T mutant GFP was found to fold four times faster than wtGFP, making it useful for applications requiring rapid detection of fluorescence following protein production. The faster folding may also partially explain the increased emission levels from the mutant protein. Additional mutagenesis of primary GFP mutants yielded versions of GFP with multiple mutations and enhanced emission properties. Two variants had lower wavelength emission than wtGFP, making it possible to obtain three distinct color emissions. Fluorescence resonance energy transfer (FRET) between two version of GFP was also demonstrated.21 In another study, optimized combinatorial mutagenesis of amino acid residues 64-69 was used to create mutations in the GFP chromophore.22 The mutant proteins were screened for GFP with an enhanced excitation maximum using Digital Imaging Spectroscopy. These mutant proteins were named “Red-Shifted” GFP (RSGFP, Table 7.1). No comparison of their fluorescence intensity with wtGFP was carried out. Two other GFP mutants (GFP/2031

Using Green Fluorescent Protein (GFP) as a Biomarker or Bioreporter for Bacteria

103

Table 7.1. GFP mutants. List of GFP variants giving the mutant name, amino acid changes, excitation wavelength, emission wavelength, and relative intensity to wtGFP GFP name

Residues mutatated

Excitation (nm)a

Emission (nm)b

Relative intensityc

wtGFP

None

395/470

509

1

2

H9

S202F

398

511

1.17

16

P9

I167V

471/396

502

1.66

16

P11

I167T

471/396

502

1.88

16

P4

Y66H

382

448

0.57

16

W

Y66W

458

480

ND

16

S65-T

S65T

490

510

6

20

Cycle 3

F100S, M145T, V164A

385

510

18

25

RSGFP4

F64M, S65G, Q69L

490

505

ND

22

GFPmut1 (EGFP)

F64L, S65T

488

507

30-50

24

GFPmut2

S65A, V68L, S72A

481

507

80-100

24

GFPmut3

S65G, S72A

501

511

70-80

24

GFPT203I

T203I

400

512

ND

23

GFPE222G

E222G

481

506

ND

23

GFPmut3* gfp(AAV, LVA, LAA, or ASV)

S2R, S65G,S72A GFPmut3* with instability tail added

501 501

511 511

70-80 70-80

26 26

S65A

S65A

ND

505

2

70

S65G

S65G

ND

513

1

70

P4-3

Y66H, Y145F

381

445

1

21

W7

Y66W, N146I, M153T, V163A, N212K

433

475

1

21

V2

Y66W, I123V, Y145H, H148R, M153T, V163A, N212K

432

480

1

21

P4-1

S65T, M153A, K238E

504

514

ND

21

ND–no data was given in the paper sited aDifferent methods were used to determine peak excitation. See the reference for details. bDifferent methods were used to determine peak emission. See the reference for details. cDifferent methods were used to determine relative intensity. See the reference for details.

Ref.

104


and GFP222G, Table 7.1) were created using a strain of E. coli deficient in DNA polymerase proofreading.23 One mutant was not fluorescent when excited with 395 nm light; the other mutant was not fluorescent when excited with 470 nm light, although they were fluorescent when excited at 400 or 481 nm, respectively. Neither of the corresponding mutations mapped to the chromophore. In another study, a codon-based mutagenesis scheme was used to mutate amino acid residues in ositions 55-74 and a FACS machine to isolate red-shifted GFP mutants, mut1, mut2 and mut3 (Table 7.1) with increased fluorescence intensity (30-100x wtGFP).24 These authors found that 90-100% of the mutant protein was soluble, which probably contributes to the enhanced level of emission. The authors also suggested that the mutations may result in faster chromophore formation and increased 488 nm excitation. Using DNA shuffling, a non-redshift mutant of GFP was isolated that remains soluble and has emission levels 18 times higher that of wtGFP.25 The different mutants described above have greatly increased the utility of GFP as a biomarker/bioreporter in bacterial systems. Variants that change the stability of GFP in bacteria have also been developed. Andersen et al added short peptide sequences to GFP that cause bacterial proteases to degrade the protein at different rates, generating GFP variants with shorter half lives.26 These mutant GFPs allow studies of temporal gene expression. The Gfp gene has also been specifically mutated to optimize expression in non-bacterial systems. Many of these mutants have codon usage optimized for a particular organism. For example, GFP mutants have been developed for yeast, plants and mammalian cells (For reviews see refs. 3,27). Therefore, it is now possible for researchers to select from a number of GFP variants with different excitation and emission maxima and fluorescence levels (see Table 7.1). Careful selection of a GFP variant suited for a particular application can maximize the utility of this marker.28 In some cases, there are problems that remain to be overcome, even when using the more optimized mutant derivatives of GFP. For example, clonal populations of bacteria expressing mutant derivatives of GFP may sometimes still exhibit variable levels of fluorescence.29 There have also been reports of no detectable fluorescence in bacteria containing GFP, even when the Gfp gene is controlled by a promoter known to be active in that bacterium. For example, Kremer et al observed blue color in all E. coli cells harboring a plasmid with the lacZ gene controlled by a heat shock promoter (hsp60).30 However, when the lacZ gene in this plasmid was replaced with the Gfp gene, no fluorescence was observed. Egener et al. (1998) observed fluorescence in Azoarcus sp BH72 harboring the gene encoding GFPmut2 under the control of the nifH promoter.31 However, they could not detect fluorescence in bacteria harboring the genes encoding wtGFP or the P11 mutant GFP expressed from the same promoter. Bacteria harboring a vector containing a kanamycin resistance gene and the P11 Gfp gene, both expressed from a PpsbA promoter, were resistant to kanamycin but were not fluorescent (Stoltzfus and de Bruijn, unpublished data). The exact reasons for lack of detectable fluorescence have not been elucidated. Toxicity of GFP can also be a problem when GFP is expressed at high levels in bacteria. For example, E. coli cells harboring high copy-number plasmids strongly expressing GFPmut1were found to be prone to cell lysis.32 However, it is now usually the case that by careful choice of mutant GFP, expression levels and copy number (for example by chromosomal integration) most if not all of these problems can be overcome.

7.4. Detection of GFP As GFP requires no additional substrate for fluorescence,2 detection is basically noninvasive, requiring no pre-detection processing and allows rapid and accurate in vivo/in


105

situ analyses. GFP also allows detection of tagged bacteria regardless of the energy status of the cells.17,33 A wide variety of detection methods suitable for analysis of bacteria expressing GFP are available which enables optimization of the experimental design to address multiple relevant biological questions without the compromises sometimes associated with biomarkers that require special detection protocols. These methods include the following: 1. visualization of bacterial colonies on plates using blue light illumination, 2. visualization of colonies or individual cells using an epifluorescence microscope, 3. visualization of single cells and three dimensional imaging of the pattern of cell aggregation/colonization in samples using a laser-scanning confocal microscope, 4. visualization of gross aggregation/colonization patterns by tagged bacteria on samples such as plant tissue using fluorescence stereomicroscopy, 5. counting fluorescent cells using flow cytometry, 6. screening bacterial collections using fluorescent microtiter plate readers, and 7. quantitation of fluorescence in suspensions using spectrofluorimetry. In some cases the use of image enhancement software may be needed to distinguish the fluorescence of tagged bacteria from background autofluorescence, such as observed in root or soil samples.17,34 Several reviews detailing these techniques are available (for reviews see refs. 10,11, 35). An important development has been the development of a filter set optimized for detection of GFP by epifluorescence microscopy.36 More specialized techniques for observing GFP-mediated fluorescence have also been described. For example, scanning near-field optical/atomic force microscopy has been used for determination of the spatial localization of tagged bacteria in liquid,37 and video-endoscopy has been employed to detect GFP expression in situ.38 Specialized software can create three-dimensional images from optical sections made from samples containing GFP labeled bacteria using a laser scanning confocal microscope. Because GFP detection is noninvasive, real time and time-lapse videos can be used to track cells (or proteins) labeled with GFP (For a review see 5). Studies using different GFP mutants have been carried out using double and even triple labeling allowing simultaneous detection of multiple targets in the same sample.21,39-41

7.5. GFP as a Biomarker in Bacteria GFP has been used as a biomarker in a variety of bacterial strains. A partial list of the bacterial species studied and the types of studies performed is presented in Table 7.2. Use of constitutive or inducible promoters to express GFP in bacterial cells allows tracking of tagged cells in the environment and visualization of their location.10 Constitutive expression can be obtained by cloning GFP into a vector containing a promoter known to be constitutively expressed in the bacterium of interest (for example, see ref. 17). Alternatively, constitutive expression can be obtained by random insertion of a promoterless Gfp gene into the bacterial genome and screening for strains that are fluorescent under particular physiological conditions.19 Inducible expression of GFP can be obtained using inducible promoters such as Plac or PF10 .42,43 Bacteria marked with GFP can also be monitored for movement and survival in the environment. For example, a promoterless GFP-containing transposon was used to obtain fluorescent P. putida strains and to monitor their movement through columns of defined medium.19 These authors also performed similar experiments with E. coli cells harboring a plasmid containing the Gfp gene under control of the inducible Plac promoter. In addition, E. coli cells with GFP controlled by a constitutive promoter were monitored to study their survival in stream water.44 GFP was also used to monitor Moraxella cells during degradation of p-nitrophenol in broth cultures and soil by enumeration of green fluorescent colonies.45 Tombolini et al used a Tn5 derivative transposon containing the Gfp gene expressed

Type of study

Localize proteins involved in septum formation and study their interactions

Create promoter fusions

Monitor cells during bioremediation of 4-chlorophenol contaminated soil

Observe Azoarcus expressing nif genes in soil and on rice roots

Localize proteins involved in sporulation septum formation


Localize proteins involved in septum formation

Localize proteins involved in chromosomal partitioning

Localize proteins to determine where processing takes place

Localize Hbsu

Localize gene expression and proteins involved in sporulation

Localize proteins involved in chromosomal partitioning

Organism

Agrobacterium tumefaciens A136

Alcaligenes eutrophus

Arthrobacter chlorophenolicus A6

Azoarcus sp BH72

Bacillus megaterium

Bacillus subtilis

Bacillus subtilis

Bacillus subtilis

Bacillus subtilis

Bacillus subtilis

Bacillus subtilis

Bacillus subtilis

chromosomal integration









mini-Tn5

mini-Tn5

plasmid

Type of Vector

pDL50B

pSG1141,pSG1147

pPK9C8

pF1

pSG1517

pSG1044

pCW28

pIIE-GFP-T

pBHFN35

pUTgfp

pAG408

pAEG3, pZEG3, and related plasmids

Vector Name

S65-T

wtGFP

wtGFP

S65-T

S65-T

wtGFP

wtGFP

GFPmut2

P11

Cycle 3

GFPmut2

GFP mutant

64

72

63

60

62

59

58

57

31

46

68

56

Ref.

Table 7.2. GFP Marked Bacteria. List of bacteria in which GFP has been used as a biomarker/bioreporter. The table lists the genus and species of bacteria used in the study, the type of vector, the name of the vector and the GFP variant carried on the vector

106 Tracking Genetically-Engineered Microorganisms

plasmid plasmid

Study chromosome partitioning




Bacillus subtilis

Bacillus subtilis

Escherichia coli

pGFP pUTgfplux

plasmid mini-Tn5

Monitor floc stability in activated sludge

Detection and counting of bacteria using GFP and monitoring their energy status using luxAB

plasmid

Study chromosome and plasmid partitioning

Escherichia coli K12

Monitor cells during degradation of p.-nitrophenol in broth and in soil

Observe infection of macrophages and epithelial cells

Moraxella sp.

Mycobacterium avium

Escherichia coli K12 Study plasmid partitioning HB101 Escherichia coli NM522 Track movement of bacteria through a column of sand

plasmid

Monitor persistence of bacteria in stream water using microscopy

Escherichia coli JM109

plasmid

pWES4

pJBA28

pGFP

plasmid mini-Tn5

pSopB-GFP

pSg20

pGFP


pURE-RD-GFP

plasmid

plasmid


Escherichia coli JM105

plasmid

Correlate ureD expression with urea concentrations

Escherichia coli DH5a

pK3G

pDR112,pDR123

Escherichia coli 01 Escherichia coli DH5a

plasmid



Escherichia coli

pAG, pCSK100, pGZ,pZG

pCH50

pSG1902

pSG1151

pAT3, pAT11, pCW13, pCW15, pCW33, pCW37

Vector Name

Escherichia coli Escherichia coli

phage




Study the expression of genes regulated by sporulation

Bacillus subtilis

Type of Vector

Type of study

Organism

wtGFP

GFP mut2

wtGFP

S65-T

wtGFP

GFPmut2

GFPmut2

P11

wtGFP

GFPmut2

GFPmut2

GFPmut2

S65-T

GFPmut1

wtGFP

GFP mutant

52

45

19

67

66

44

56

77

33

18

76

75

55

74

61

65

73

Ref.

Using Green Fluorescent Protein (GFP) as a Biomarker or Bioreporter for Bacteria 107

Observe ureD gene expression by bacteria in bladders and kidneys of infected mice

Proteus mirabilis HI4320 Pseudoaltermonas sp. plasmid

plasmid

plasmid

plasmid

plasmid

plasmid mini-Tn5

Pseudomonas aeruginosa Create promoter fusions PA01

Pseudomonas chlororaphis MA 342

Visualize pattern of cell aggregation on barley seeds

plasmid

Pseudomonas aeruginosa Observe fluorescent intensity and plasmid stability and distinguish tagged bacteria in mixed cultures

Investigate colonization behaviour of a marine mini-Tn10 isolate on squid pen (a natural biodegradable substrate)

Use as a promoter probe for bacterial promoters

Study expression patterns of different promoters in bacteria grown in macrophages

Mycobacterium marinum Mycobacterium smegmatis

Mycobacterium smegmatis

Visualize bacteria infecting host cells and count infected cells using flow cytometry

Mycobacterium bovis BCG

Test for fluorescence

plasmid

Observe infection of cell line from mice


Mycobacterium smegmatis

Used to test drug susceptibility, response to plasmid environmental signals and localize bacteria in vivo

plasmid

Study expression patterns of different promoters in bacteria grown in macrophages



Observe infection of macrophages and epithelial cells

Mycobacterium avium

Type of Vector

Type of study

Organism wtGFP

GFP mutant

PUTgfp2

pIVET-GFP

pSMC2

mini-Tn10-gfp-kan

pURE-RD-GFP

pGFM-12

P11

GFPmut3

GFPmut2

GFPmut2

GFPmut2

wtGFP

wtGFP

wtGFP

phsp60-gfp, pMlaphc-gfp, pMtaphc-gfp, pmtra-gfp, ptbprc3-gfp pGFM-11

wtGFP

S65-T

wtGFP

pFPV2

pYL mutGFP

pGFM-11

phsp60-gfp, wtGFP pMlaphc-gfp, pMtaphc-gfp, pmtra-gfp, ptbprc3-gfp

pMV306(hsp 60/gfp)

Vector Name

49

69

50

80

77

30

30

78

79

51

30

78

52

Ref.

108 Tracking Genetically-Engineered Microorganisms

Counting of bacteria using GFP and monitoring their energy status using luxAB

Visualize bacteria infecting host cells and count infected cells using flow cytometry

plasmid

pFPV1

TOL-gfp (RP1::GFP)

plasmid

Yersinia pseudotuberculosis

pFVP25

plasmid

Salmonella typhimurium Find acid induced promoters and test for expression by bacteria in macrophages Serratia marcescen Monitor floc stability in activated sludge

pFPV1

plasmid

pUTgfp

Detect VBNC cells in groundwater microcosms

Salmonella typhi

Salmonella typhimurium Visualize bacteria infecting host cells and count infected cells using flow cytometry

mini-Tn5


Sinorhizobium meliloti MB501

pTB93F

pAG408


plasmid plasmid

mini-Tn5

TOL-gfp (RP1::GFP)

plasmid

Visualize nodulation of Meticago sativa by bacteria

pJBA30, pJBA26

mini-Tn5


Tn5GFP1

pAG408

pRL765gfp

Tn5 derivative

mini-Tn5

Tn5

Pseudomonas sp B13 Sinorhizobium meliloti 1021

Pseudomonas Track movement of bacteria through putida mt-2 a column of sand Pseudomonas putida R1 Observe gene induction and bacterial location in a bioreactor Pseudomonas Monitor plasmid transfer by conjugation putida KT2442


pUTgfplux

mini-Tn5

Monitor conjugal plasmid transfer in the phylloplane of bush bean

Pseudomonas fluorescens AS12 Pseudomonas fluorescens SBW25

Pseudomonas putida KT2442

TOL::gfpmut3b

plasmid

Monitor bacteria in culture, soil samples, and on roots with flow cytometry and microscopy

pGB5

Pseudomonas fluorescens A506

plasmid

Observe fluorescent intensity and plasmid stability and localize bacteria on plant roots

Vector Name

Pseudomonas fluorescens WCS365

Type of Vector

Type of study

Organism

wtGFP

wtGFP

GFPmut3

wtGFP

wt GFP

GFPmut2

S65-T

Cycle 3

tGFP

GFPmut3b

wtGFP

Cycle 3

P11

GFPmut3b

P11

GFPmut2

GFP mutant

79

18

79

79

47

56

34

68

42

71

19

68

33

43

17

50

Ref.

Using Green Fluorescent Protein (GFP) as a Biomarker or Bioreporter for Bacteria 109

110


from a constitutive promoter to tag P. fluorescens cells, and showed that the tagged cells could be detected by flow cytometry, epifluorescent microcopy and laser-scanning confocal microscopy. 17 In addition, dual tagging experiments using GFP and bacterial luciferase has allowed simultaneous tracking of tagged bacteria cells and monitoring of their energy status (by luciferase activity) in soil microcosms.33 Similarly, Arthrobacter chlorophenolicus cells were tagged with either GFP or the firefly luciferase gene (luc) in order to monitor Arthrobacter cells during bioremediation of 4-chlorophenol in soil.46 GFP-tagged cells could be enumerated in soil by flow cytometry, after extraction of the bacterial soil fraction by density gradient centrifugation to remove soil particles and debris that would otherwise interfere with flow cytometric measurements.33,46 GFP fluorescent A. chlorophenolicus 46 or P. fluorescens 33 cells could be counted at relatively stable levels after inoculation to soil, compared to counts based on CFU or luciferase expression, which decreased during incubation. These experiments demonstrate the utility of GFP as a stable marker for counting the total number of tagged cells in environmental samples, regardless of their metabolic activity or culturability. However, it is possible that some of the cells that were counted in such experiments are dead.33,46 Recent studies in our laboratory and in others have shown that viable but nonculturable cells (see Chapter 1) can still retain GFP fluorescence.47,48 GFP can also be used to monitor transfer of plasmids between bacteria. Christensen et al used a plasmid containing GFP controlled by an inducible promoter, PF10, to monitor plasmid transfer on semi-solid surfaces.42 The P. putida donor strain did not contain the T7 RNA polymerase gene and therefore the T7 PF10 promoter was inactive and the bacteria were not fluorescent. The recipient strain carried the T7 RNA polymerase gene integrated in its chromosome. T7 RNA polymerase activates the PF10 promoter and hence any recipient that captured PF10-GFP became fluorescent. Thus, fluorescence resulted only when the plasmid was transferred to the recipient strain. A similar system was used with GFP expression regulated by Plac to observe plasmid transfer in the phylloplane of bush beans and on polycarbonate filters.43

7.5.1. Localizing Bacteria in situ As illustrated above, bacteria marked with GFP can clearly be visualized in situ, allowing identification of the environmental niche they occupy. One research area that has greatly benefited from the use of GFP has been the localization of plant-associated bacteria. An example of the use of GFP as a marker to localize specific bacteria directly on plant surfaces is shown in Fig. 7.1. In this study, the biocontrol strain P. chlororaphis MA 342, which controls fungal disease on cereal crops, was chromosomally tagged with two copies of the Gfp gene to enhance GFP fluorescence, and enabling visualization of the pattern of colonization of the cells on barley seeds by fluorescence stereomicroscopy and confocal microscopy (Fig. 7.1).49 We have also used bacteria marked with a transposon containing GFP to monitor the interactions between rice roots and endophytic bacteria (Stoltzfus and de Bruijn, unpublished data). Moreover, Gage et al used confocal microscopy to monitor early events in rhizobial infection and nodule formation on alfalfa using a Sinorhizobium meliloti isolate carrying a plasmid constitutively expressing GFP,34 and Bloemberg et. al used a combination of phase contrast and fluorescence microscopy to distinguish Pseudomonas aeruginosa cells containing plasmid-borne GFP from untagged Burkholderia cepacia bacteria attached to an abiotic surface.50 These authors also monitored GFP-tagged P. fluorescens bacteria associated with the roots of tomato seedlings by fluorescence microscopy. There are several other published examples of the use of GFP as a marker for in situ localization of bacteria in other kinds of environments. For example, GFP-marked E. coli and Serratia marcescens cells have also been used to study flocculation in activated sludge by


111

Fig. 7.1. Confocal microscope projection of a stack of images of individual GFP-tagged Pseudomonas chlororaphis MA342G2 cells (white spots on this figure) distributed on the outer surface of a barley seed coat or glume (large cells). A model LSM501 laser scanning confocal microscope (Carl Zeiss, Jena, Germany) was used. The images are the result of pseudocolor merging of the output of three channels. Three-dimensional rendering of the stack of images was obtained by using the software 3D for LSM510, version 1.4 (Carl Zeiss).

confocal microscopy.18 In addition, Mycobacterium bovis cells were marked with GFP to follow their infection of mice cells using flow cytometry.30,51 The infection of human macrophages and epithelial cells by Mycobacterium avium marked with GFP has also been observed,52 and Valdivia et al used GFP to visualize infection of live mammalian cells by Salmonella typhimurium, Yersinia pseudotuberculosis and Mycobacterium marinum.53

7.6. GFP as a Bioreporter in Bacteria The fusion of GFP to a protein of interest and monitoring of the expression of the fusion product in vivo permits the assessment of the cellular localization of the fusion protein (For reviews see refs. 3,5,6,8). Although GFP looses its fluorescence when truncated,3 the full-length protein can be used successfully in both C-terminal and N-terminal protein fusions.54 GFP has been used extensively as a bioreporter to study cell division in bacteria. For example, Ma et al have created fusions of the cell division proteins FtsZ and FtsA to GFP in E. coli cells and reported that both C-terminal and N-terminal fusions of FtsZ with GFP showed midcell localization and polymerization.55 FtsZ and FtsA interactions have also been examined with GFP fusions in Agrobacterium tumefaciens and S. meliloti.56 GFP has also been used to study proteins involved in septation during vegetative division and sporulation in Bacillus.57-61

112


The localization of proteins involved in chromosome behavior has also been studied using GFP. 62-64 Webb et al used a GFP-LacI fusion to localize specific regions of the B. subtilis chromosome in vegetative and sporulating cells by inserting a lacO cassette at specific sites in the chromosome.65 The same system was used to compare chromosome and plasmid behavior during replication of E. coli.66,67 Promoterless GFP constructs can also be used to screen for promoters that are induced under a specific set of physiological conditions. For example, Kremer et al inserted fragments of the M. bovis genome in front of the Gfp gene and found that 3 to 5% of the chimeric genes showed differing levels of fluorescence on agar plates.30 Suarez et al. (1997) developed both plasmid and mini-Tn5-based promoterless GFP vectors.68 P. putida, Pseudomonas sp B13 and Alcaligenes eutrophus were mutated with mini-Tn5::GFP. Up to 5% of the bacteria with chromosomal integration of the mini-Tn5 were fluorescent, displaying varied levels of expression. Moreover, a promoterless GFP vector was developed that allows rapid selection for active promoters by fusing aspartate β-semialdehyde dehydrogenase with GFP and using it as a selectable marker in Asd- strains.69 Another example of the utility of GFP as a bioreporter is the construction of pGreenscript.70 In this vector GFP replaces the β-galactosidase (lacZ gene) as the marker for insertion of DNA in the multiple cloning site of the popular cloning vector pBluescript. Biomarking can be combined with bioreporting to obtain spatial and temporal information on gene expression in situ. For example, a nifH promoter-GFP fusion has been used to monitor expression of nitrogen-fixing genes in Azoarcus sp. BH72 in soil and on rice roots.31 Moreover, Møller et al used P. putida cells containing the Gfp gene expressed from promoters controlling the expression of proteins involved in the biodegradation of toluene to visualize gene expression in flow chambers and observed the effects of various community members on gene induction.71 These studies demonstrate that key activities of microbial communities in environmental samples can be specifically investigated in situ using GFP as a reporter.

7.7. Conclusion The large number of studies employing GFP as a biomarker/bioreporter clearly demonstrate the value and versatility of GFP. GFP is particularly useful for visualization of single cells or cell aggregates in environmental samples without the need for any substrate addition. Cells tagged with GFP can be monitored by a range of fluorescence detection methods including flow cytometry and fluorescence microscopy. GFP also can be used to quantitate the number of cells of a particular cell population, regardless of the energy status of the cells. This is in contrast to most of the other existing markers available, such as luciferase markers, that require the cells to be metabolically active to be detected. While problems with GFP as a biomarker/bioreporter in bacteria have been encountered, many have been overcome by using mutant versions of GFP, adding appropriate translation initiation sites, or adding flexible linkers to protein fusions. The many successful studies using GFP and the continued development of new techniques incorporating GFP will undoubtedly lead to the use of GFP in more species of bacteria to answer diverse questions in molecular microbial ecology.

Acknowledgments Jon R. Stoltzfus and Frans J. de Bruijn gratefully acknowledge support for studies carried out at MSU from the NSF Center for Microbial Ecology (CME; grant no. DEB9120006), from the International Rice Research Institute (IRRI) and from the Department of Energy (DOE; grant no. DE-FG02-91ER20021). Janet K. Jansson gratefully acknowledges support from the Swedish Council for Engineering Science, the Carl Tryggers Foundation the Swed-


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ish Foundation for Environmental Research and the Swedish Foundation for Strategic Research. Both F.J. de Bruijn and J.K. Jansson are partners of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Prasher DC, Eckenrode VK, Ward WW et al. Primary structure of the Aequorea victoria green-fluorescent protein. Gene 1992; 111(2):229-233. 2. Chalfie M, Tu Y, Euskirchen G et al. Green fluorescent protein as a marker for gene expression. Science 1994; 263(5148):802-805. 3. Cubitt AB, Heim R, Adams SR et al. Understanding, improving and using green fluorescent proteins. Trends Biochem Sci 1996; 20(11):448-455. 4. Errampalli D, Leung, K, Cassidy MB et al. Applications of the green fluorescent protein as a molecular marker in environmental microorganisms. J Microbiol Meth 1999; 35:187-199. 5. Gerdes HH and Kaether C. Green fluorescent protein: applications in cell biology. FEBS Lett 1996; 389(1):44-47. 6. Jansson JK, and de Bruijn FJ. Biomarkers and Bioreporters to track microbes and monitor their gene expression. In: Davies J, ed. Manual of Industrial Microbiology and Biotechnology. 2nd ed. Washington DC: ASM Press, 1999:651-665. 7. Kendall JM, Badminton MN. Aequorea victoria bioluminescence moves into an exciting new era. TIBTECH 1998; 16:216-224. 8. Misteli T and Spector DL. Applications of the green fluorescent protein in cell biology and biotechnology. Nature Biotechnology 1997; 15(10):961-964. 9. Tsien RY. The green fluorescent protein. Ann Rev Biochem 1998; 67:509-544. 10. Unge A, Tombolini R, Davey ME et al. GFP as a Marker Gene. In: Akkermans ADL, van Elsas JD, de Bruijn FJ, eds. Molecular Microbial Ecology Manual. Dordrecht, The Netherlands: Kluwer Academic Publishers, 1998; 6.1.13:1-16. 11. Tombolini R and Jansson JK Monitoring of GFP tagged bacterial cells. Methods in Molecular Biology 1998; 102:285-298. 12. Prasher DC. Using GFP to see the light. Trends Genet 1995; 11(8):320-323. 13. Cody CW, Prasher DC, Westler WM et al. Chemical structure of the hexapeptide chromophore of the Aequorea green-fluorescent protein. Biochemistry 1993; 32(5):1212-1218. 14. Morise JG, Shimomura O, Johnson FH et al. Intermolecular energy transfer in the bioluminescent system of Aequorea. Biochemistry 1974; 13:2656-2662. 15. Ward WW, Cody CW, Prasher DC et al. Spectrophotometric identity of the energy-transfer chemophores in Renilla and Aequorea green-fluorescent protein. Photochem Photobiol 1980; 31:611-615. 16. Heim R, Prasher DC, and Tsien RY. Wavelength mutations and posttranslational autoxidation of the green fluorescent protein. Proc Natl Acad Sci 1994; 91(26):12501-12504. 17. Tombolini R, Unge A, Davey MA et al. Flow cytometric and microscopic analysis of GFPtagged Pseudomonas fluorescens bacteria. FEMS Microbiol Ecol 1997; 22:17-28. 18. Olofsson AC, Zita A and Hermansson M. Floc stability and adhesion of green-fluorescentprotein-marked bacteria to flocs in activated sludge. Microbiology 1998; 144(2):519-528. 19. Burlage RS, Yang ZK, and Mehlhorn T. A transposon for green fluorescent protein transcriptional fusions: Application for bacterial transport experiments. Gene 1996; 173(1);53-58. 20. Heim R, Cubitt AB and Tsien RY. Improved green fluorescence. Nature 1995; 373(6516): 663-664. 21. Heim R, and Tsien RY. Engineering green fluorescent protein for improved brightness, longer wavelength and fluorescence resonance energy transfer. Curr. Biol. 1996; 6(2):178-182. 22. Delagrave S, Hawtin RE, Silva CM et al. Red-shifted excitation mutants of the green fluorescent protein. Biotechnology 1995; 13(2):151-154. 23. Ehrig T, O’Kane DJ and Prendergast FG. Green-fluorescent protein mutants with altered fluorescence excitation spectra. FEBS Lett. 1995; 367(2):163-166.

114


24. Cormack BP, Valdivia RH and Falkow S. FACS-optimized mutants of the green fluorescent protein (GFP). Gene 1996; 173(1):33-38. 25. Crameri A, Whitehorn EA, Tate E et al. Improved green fluorescent protein by molecular evolution using DNA shuffling. Nat Biotechnol 1996; 14(3):315-319. 26. Andersen JB, Sternberg C, Poulsen LK et al. New unstable variants of green fluorescent protein for studies of transient gene expression in bacteria. Appl Environ Microbiol 1988; 64(6):2240-2246. 27. Stauber RH, Horie K, Carney P et al. Development and applications of enhanced green fluorescent protein mutants. Biotechniques 1998; 24(3):462-466. 28. Patterson GH, Knobel SM, Sharif WD et al. Use of the green fluorescent protein and its mutants in quantitative fluorescence microscopy. Biophys J 1997; 73(5):2782-2790. 29. Matthysse AG, Stretton S, Dandie C et al. Construction of GFP vectors for use in Gramnegative bacteria other than Escherichia coli. FEMS Microb Lett 1996; 145(1):87-94. 30. Kremer L, Baulard A, Estaquier J et al. Green fluorescent protein as a new expression marker in mycobacteria. Molecular Microbiology 1995; 17(5):913-922. 31. Egener T, Hurek T and Reinhold-Hurek B. Use of green fluorescent protein to detect expression of nif genes in Azoarcus sp. BH72, a grass-associated diazotroph, on rice roots. Molecular Plant-Microbe Interactions 1998; 11(1):71-75. 32. Miller WB and Lindow SE An improved GFP cloning cassette designed for prokaryotic transcriptional fusions. Gene 1997; 191(2):149-153. 33. Unge A, Tombolini R, Molbak L et al. Simultaneous monitoring of cell number and metabolic activity of specific bacterial populations with a dual gfp/luxAB marker system. Appl Environ Microbiol 1999; 65(2):813-821. 34. Gage DJ, Bobo T and Long SR. Use of green fluorescent protein to visualize the early events of symbiosis between Rhizobium meliloti and alfalfa (Medicago sativa). J Bacteriol 1996; 178(24):7159-7166. 35. Khodjakov A, Cole RW and Rieder CL. A synergy of technologies: Combining laser microsurgery with green fluorescent protein tagging. Cell Motil Cytoskeleton 1997; 38(4):311-317. 36. Zylka MJ and Schnapp BJ. Optimized filter set and viewing conditions for S65T mutant of GFP in living cells. Biotechniques 1996; 21(2):220-226. 37. Tamiya E, Iwabuchi S, Nagatani N et al. Simultaneous topographic and fluorescence imagings of recombinant bacterial cells containing a green fluorescent protein gene detected by a scanning near-field optical/atomic force microscope. Anal Chem 1997; 69(18):3697-3701. 38. Flotte TR, Beck SE, Chesnut K et al. A fluorescence video-endoscopy technique for detection of gene transfer and expression. Gene Ther 1998; 5(2):166-177. 39. Yang TT, Cheng L and Kain SR. Optimized codon usage and chromophore mutations provide enhanced sensitivity with the green fluorescent protein. Nucleic Acids Res 1996; 24(22):4592-4593. 40. Yang TT, Sinai P, Green G et al. Improved fluorescence and dual color detection with enhanced blue and green variants of the green fluorescent protein. J Biol Chem 1998; 273(14):8212-8216. 41. Lybarger L, Dempsey D, Patterson GH et al. Dual-color flow cytometric detection of fluorescent proteins using single-laser (488-nm) excitation. Cytometery 1998; 31(3):147-152. 42. Christensen BB, Sternberg C, and Molin S. Bacterial plasmid conjugation on semi-solid surfaces monitored with the green fluorescent protein (GFP) from Aequorea victoria as a marker. Gene 1996; 173(1):59-65. 43. Normander B, Christensen BB, Molin S et al. Effect of bacterial distribution and activity on conjugal gene transfer on the phylloplane of the bush bean. Appl Environ Microbiol 1998; 64(5):1902-1909. 44. Leff LG and Leff AA. Use of green fluorescent protein to monitor survival of genetically engineered bacteria in aquatic environments. Appl Environ Microbiol 1996; 62(9):3486-3488. 45. Tresse O, Errampalli D, Kostrzynska M et al. Green fluorescent protein as a visual marker in a p-nitrophenol degrading Moraxella sp. FEMS Microbiol Lett 1998; 164:187-193.


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46. Elväng AM, Westerberg K, Jernberg C et al. Monitoring bioremediation of 4-chlorophenol-contaminated soil using gfp or luc-tagged Arthrobacter chlorophenolicus A6 cells. Submitted. 47. Cho J-C, Kim S-J. Green fluorescent protein-based direct viable count to verify a viable but non-culturable state of Salmonella typhi in environmental samples. J Microbiol Meth 1999; 36:227-235. 48. Lowder M, Unge A, Jansson JK et al. The effect of starvation and the VBNC state on GFP fluorescence in Pseudomonas fluorescens. Submitted 49. Tombolini R, Van der Gaag DJ, Gerhardson, B et al. Colonization pattern of the biocontrol strain Pseudomonas chlororaphis MA 342 on barley seeds visualized by using green fluorescent protein. Appl Environ Microbiol 1999; 65:3674-3680. 50. Bloemberg GV, O’Toole GA, Lugtenberg BJJ et al. Green fluorescent protein as a marker for Pseudomonas spp. Appl Environ Microbiol 1997; 63(11):4543-4551. 51. Luo Y, Szilvasi A, Chen X et al. A novel method for monitoring Mycobacterium bovis BCG trafficking with recombinant BCG expressing green fluorescent protein. Clin Diagn Lab Immunol 1996; 3(6):761-768. 52. Parker AE and Bermudez LE. Expression of the green fluorescent protein (GFP) in Mycobacterium avium as a tool to study the interaction between Mycobacteria and host cells. Microb Pathog 1997; 22(4):193-198. 53. Valdivia RH, Hromockyj AE, Monack D et al. Applications for green fluorescent protein (GFP) in the study of host-pathogen interactions. Gene 1996; 173(1):47-52. 54. Wang S and Hazelrigg T. Implications for bcd mRNA localization from spatial distribution of exu protein in Drosophila oogenesis. Nature 1994; 369(6479):400-403. 55. Ma X, Ehrhardt DW and Margolin W. Colocalization of cell division proteins FtsZ and FtsA to cytoskeletal structures in living Escherichia coli cells by using green fluorescent protein. Proc Natl Acad Sci 1996; 93(23):12998-13003. 56. Ma X, Sun Q, Wang R et al. Interactions between heterologous FtsA and FtsZ proteins at the FtsZ ring. J Bacteriol 1997; 179(21):6788-6797. 57. Barak I, Behari J, Olmedo G et al. Structure and function of the Bacillus SpoIIE protein and its localization to sites of sporulation septum assembly. Mol Microbiol 1996; 19(5):1047-1060. 58. Arigoni F, Pogliano K, Webb CD et al. Localization of protein implicated in establishment of cell type to sites of asymmetric division. Science 1995; 270(5236):637-640. 59. Edwards DH and Errington J. The Bacillus subtilis DivIVA protein targets to the division septum and controls the site specificity of cell division. Mol Microbiol 1997; 24(5):905-915. 60. Ju J, Luo T and Haldenwang WG. Bacillus subtilis pro-sigmaE fusion protein localizes to the forespore septum and fails to be processed when synthesized in the forespore. J Bacteriol 1997; 179(15):4888-4893. 61. Wu LJ, Feucht A and Errington J. Prespore-specific gene expression in Bacillus subtilis is driven by sequestration of SpoIIE phosphatase to the prespore side of the asymmetric septum. Genes Dev 1998; 12(9):1371-1380. 62. Glaser P, Sharpe ME, Raether B et al. Dynamic, mitotic-like behavior of a bacterial protein required for accurate chromosome partitioning. Genes Dev 1997; 11(9):1160-1168. 63. Kohler P and Marahiel MA. Association of the histone-like protein HBsu with the nucleoid of Bacillus subtilis. J Bacteriol 1997; 179(6):2060-2064. 64. Lin DCH, Levin PA and Grossman AD. Bipolar localization of a chromosome partition protein in Bacillus subtilis. Proc Natl Acad Sci 1997; 94(9):4721-4726. 65. Webb CD, Teleman A, Gordon S et al. Bipolar localization of the replication origin regions of chromosomes in vegetative and sporulating cells of B. subtilis. Cell 1997; 88(5):667-674. 66. Gordon GS, Sitnikov D, Webb CD et al. Chromosome and low copy plasmid segregation in E. coli: visual evidence for distinct mechanisms. Cell 1997; 90(6):1113-1121. 67. Kim SK and Wang JC. Localization of F plasmid SopB protein to positions near the poles of Escherichia coli cells. Proc Natl Acad Sci 1998; 95(4):1523-1527.

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68. Suarez A, Guttler A, Stratz M et al. Green fluorescent protein-based reporter systems for genetic analysis of bacteria including monocopy applications. Gene 1997; 196(1-2):69-74. 69. Handfield M, Schweizer HP, Mahan MJ et al. ASD-GFP vectors for in vivo expression technology in Pseudomonas aeruginosa and other gram-negative bacteria. Biotechniques 1998; 24(2):261-264. 70. Inouye S, Ogawa H, Yasuda K et al. A bacterial cloning vector using a mutated Aequorea green fluorescent protein as an indicator. Gene 1997; 189(2):159-162. 71. Møller S, Sternberg C, Andersen JB et al. In situ gene expression in mixed-culture biofilms: evidence of metabolic interactions between community members. Appl Environ Microbiol 1998; 64(2):721-732. 72. Lewis PJ and Errington J. Use of green fluorescent protein for detection of cell-specific gene expression and subcellular protein localization during sporulation in Bacillus subtilis. Microbiology 1996; 142(4):733-740. 73. Webb CD, Decatur A, Teleman A et al. Use of green fluorescent protein for visualization of cell-specific gene expression and subcellular protein localization during sporulation in Bacillus subtilis. J Bacteriol 1995; 177(20):5906-5911. 74. Hale CA and de Boer PA. Direct binding of FtsZ to ZipA, an essential component of the septal ring structure that mediates cell division in E. coli. Cell 1997; 88(2):175-185. 75. Raskin DM and de Boer PA. The MinE ring: An FtsZ-independent cell structure required for selection of the correct division site in E. coli. Cell 1997; 91(5):685-694. 76. Yu XC, Tran AH, Sun Q et al. Localization of cell division protein FtsK to the Escherichia coli septum and identification of a potential N-terminal targeting domain. J Bacteriol 1998; 180(5):1296-1304. 77. Zhao H, Thompson RB, Lockatell V et al. Use of green fluorescent protein to assess urease gene expression by uropathogenic Proteus mirabilis during experimental ascending urinary tract infection. Infect Immun 1998; 66(1):330-335. 78. Dhandayuthapani S, Via LE, Thomas CA et al. Green fluorescent protein as a marker for gene expression and cell biology of mycobacterial interactions with macrophages. Mol Microbiol 1995; 17(5):901-912. 79. Valdivia RH and Falkow S. Bacterial genetics by flow cytometry: Rapid isolation of Salmonella typhimurium acid-inducible promoters by differential fluorescence induction. Mol Microbiol 1996; 22(2):367-378. 80. Stretton S, Techkarnjanaruk S, McLennan AM et al. Use of green fluorescent protein to tag and investigate gene expression in marine bacteria. Appl Environ Microbiol 1998; 64:2554-2559.

CHAPTER 8

Monitoring Persistence and Risk Assessment Following the Field Release of Pseudomonas fluorescens SBW25EeZY6KX Mark J. Bailey, Tracey M. Timms-Wilson, Richard J. Ellis, Ian P. Thompson and Andrew K. Lilley

8.1. Introduction

T

he selection of Genetically Modified Microorganisms (GMMs), or other candidate bacteria, for environmental release should be carefully considered.1 This is particularly relevant when developing inocula with known functional traits, such as for biological control, Since it is likely that the most effective inocula derive from the intended target habitat.2,3 However, prior to the field release of a GMM, it remains essential to study the organism’s autecology using enclosed facilities. Of particular importance in such studies is the capacity of the GMM to persist and cause an impact, features that are dependent on fitness and competitive ability.4-6 To some extent, these factors can be predicted from knowledge of closely related, indigenous populations.7 Data from our own releases in glasshouse and field environments confirm those of previous investigations, i.e., any impacts caused by GMMs are likely to be short lived and probably no more ecologically significant8-10 than those caused by wild type bacterial inocula.11,12 It is worthwhile noting that the apparent competitiveness of the GMM used in our studies may have been exaggerated by the artificiality of glasshouse conditions.13 We observed that bacterial diversity in the phyllosphere of glasshouse plants was in most cases significantly lower than that detected in the field. The calculated Shannon diversity index (H’) for the community colonizing immature leaves of 133 day old glasshouse grown sugar beet plants was 0.04 (4 taxa) compared to 1.114 (20 taxa) for field grown plants grown for approximately the same period. This study demonstrated that the behavior of GMMs in protected environments such as glasshouses is unlikely to be representative of the environment and can only be addressed by field studies. The aims of the investigations outlined below have been to provide appropriate information for the assessment of impact and survival of GMMs in the environment. We have undertaken a number of investigations, following a stepwise approach, to confirm the suitability of the candidate bacterium Pseudomonas fluorescens SBW25, and its genetic modification.14 These studies have been divided into three phases comprised of: Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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1. prerelease studies in contained environments2,5,13 2. the field releases to sugar beet crops4,5,15 and, 3. as discussed here, postrelease monitoring.

8.2. Construction of P. fluorescens SBW25EeZY6KX and Detection and Monitoring Methods To facilitate detection, marker genes, for antibiotic resistance (aph1, kanamycin), lactose utilization (lacZY, lactose permease and β-galactosidase—also converts X-gal to a blue pigmented product) and colorimetric detection (xylE, 2,3,catechol dioxygenase, converts catechol to a soluble yellow pigmented semi-aldehyde), were selected to provide a unique set of marker phenotypes for highly sensitive plate isolation and enumeration assays.14 These marker genes were introduced, by site-directed homologous recombination, to predetermined, nonessential sites in the choromosome,16 for the construction of P. fluorescens SBW25EeZY6KX.14,17 PCR based methods were then used for the detection of the introduced genes,18 and plate isolation methods were used for detecting mobile genetic elements acquired by the GMM from the indigenous microbial community.4,18

8.3. Case Study of the Persistence of P. fluorescens SBW25EeZY6KX in Sugar beet Crops Data gathered in microcosm and field investigations confirmed the utility of P. fluorescens SBW25EeZY6KX as a competitive phytosphere colonizer of plants when introduced as a seed dressing4,5,19 or foliar spray.20,21 In open field studies, information on the survival, establishment and dissemination of GMMs in the environment was acquired during the development and maturation of deliberately inoculated field crops. Data on the spread, persistence and survival of the inocula on plants and in soil15,21 were collected. In addition studies were undertaken that demonstrated the role of phytophageous insects in the effective dispersal of leaf colonizing populations of the GMM to other plants.22 Data on the longer term persistence of introduced inocula as a viable fraction of the soil and phytosphere microbiota or gene pool need to be gathered in an ecologically relevant environment. The approach taken and studies pertinent to the long term monitoring of genetically modified inocula and their genes have not been widely discussed. However, the persistence and fate of environmentally competent bacteria is a highly relevant issue. P. fluorescens SBW25 was selected for its ability to successfully colonize plants following seed inoculation, and was isolated directly from one of the target habitats, the sugar beet phyllosphere.23 It therefore provided an ideal opportunity to observe whether a known, habitat-adapted isolate would exhibit greater fitness in its presumed natural habitat or in alternative habitats such as soil and wheat plants. The ability of the isolate to persist had been observed during glasshouse studies where the GMM colonized the rhizoplane and rhizosphere of sugar beet representing ca. 0.05% of the pseudomonad community.13 A similar correlation between the recombinant bacterial populations and the total pseudomonad community was observed in the phyllosphere. It is considered likely that the GMM colonized those niches, in developing tissue, typically occupied by other closely related fluorescent pseudomonads and is an effective competitor due to the introduction of a large inocula to the seed (> 107 cfu/seed). In contrast to the field situation, the GMM was still present in the glasshouse rhizosphere environment on sugar beet that had been maintained for 531 days after seed inoculation, demonstrating that it survives well especially when closely associated with developing leaf and root tissue (Table 8.1).

Field Released Pseudomonas fluorescens SBW25EeZY6KX

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Table 8.1. Percentage of total colony forming units, growing on tryptic soy broth agar, identified as P. fluorescens SBW25EeZY6KX isolated from the phytosphere of glasshouse grown plants Days after Sugar beet plant habitat sampled after seed inoculation with ca. 1 x 107 cfu GMM sowing

29 50 64 71 92 103 126 133 190 220 231 531

Immature leaf

Mature leaf

Rhizosphere

Rhizoplane

Root cortex

18.6 64.4 -x 79.0 – 65.0 54.6 80.7 65.3 – 38.1 94.1

– – 65.3 22.8 36.4 3.7 10.0 – – – 11.9 0.03

– 22.6 – _ – – 0.07 – – 0.01 0.30 0.02

– – – – – – – – – 0.40 – 0.05

– – – – – – – – – 11.8 – nd

Notes, nd = no GMMs detected (limit of detection 100 cfu g-1), – = not determined. Total counts estimated by serial dilution of plant tissue homogenates spread onto Trypic Soy broth agar; GMM counts confirmed Pseudomonad selective agar (Oxoid) supplemented with 100 µg ml-1 kanamycin, and 0.01% X-gal, method and table after Thompson IP, Ellis RJ, Bailey MJ. FEMS Microbiol Ecol 1995; 17:1-14.

8.4. Survival of Recombinant in Bulk Soil Under Glasshouse Conditions Seventy one days after planting GMM inoculated sugar beet seeds in nonsterile soils in the glasshouse, bulk soil samples were collected 0-1 cm, 1-2 cm, 2-3 cm and 5-6 cm away from the root. GMM bacteria were only detected in the rhizosphere zone, < 1 cm from the root (1.06 x 104 cfu g-1) against a background total bacterial count of 9.98 x 106 cfu g-1. In addition, nonrhizosphere soil (bulk soil) was inoculated with GMM at ca 3 x 105 cfu g-1 and maintained at a moisture content approximate to field conditions (30%). The GMM survived poorly in soil, particularly at elevated temperatures or in the absence of root material. Total bacterial counts remained relatively constant irrespective of the alteration in conditions and exposure to different temperatures, 4°C, 15°C or 28°C. Field soil temperatures, at a depth of 10-20 cm, were recorded at 16°C (± 1°C) throughout the growing season. GMM numbers declined rapidly when the inoculated soil was maintained at 28°C, GMM were no longer detected after 34 days. Similarly, treatment at 15°C reduced GMM survival, as GMMs were not detected after 81 days. At 4°C a more gradual decline in GMM numbers was observed. Temperature, therefore, affects recombinant survival in the soil. These data allowed the prediction that after successful phytosphere colonization from seed inoculation in the absence of viable plant material, and a period of over-wintering, that the field-released GMM, P. fluorescens SBW25EeZY6KX, would not persist. However, despite careful design, glasshouse conditions remain artificial and therefore controlled investigations under true field conditions are necessary to determine whether inocula survived in the soil or on plant debris postharvest.

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8.5. Field Sites, Treatment and Sampling of Plants and Soil

The GMM was applied as a seed dressing (7 x 106 cfu seed-1) to commercially pelleted sugar beet (Beta vulgaris var. Amethyst) in April 1993.15 The field plots were planted in 10 by 10 row arrays surrounded by three rows of untreated plants in Evesham series clay soil at the University Farm, Wytham, UK. Nine individual 5 m2 plots of sugar beet were established. Following a Latin square design three plots were planted with untreated seeds, three with seeds inoculated with wild type P. fluorescens SBW25, and the remaining plots planted with seed inoculated with P. fluorescens SBW25EeZY6KX. The survival and persistence of the organism, together with its dispersal, were monitored until the crop was harvested in January 1994. A number of plants were left in each plot so that the persistence of the GMM on over-wintered sugar beet could be determined. Triplicate samples of soil and weed plants were also collected on a regular basis. Leaves were removed from over-wintering sugar beet plants in early March and were sampled further as new tissue developed. In April 1994 and April 1995, untreated sugar beet seeds were sown in half the area of each of the plots and monitored for colonization by the GMM. The presence of the GMM on the leaves and roots of indigenous volunteer weed species was also investigated. Weed plant samples which included Creeping Buttercup (Ranunculus repens L.), Fat hen (Chenopodium album L.), Knot grass (Polygonum avicular L.) and Thistle (Cardus teniflorus Curt.) were excavated from the ground (Table 8.2). Root and leaf samples were examined for the presence of recombinants as described for sugar beet samples. The GMM persisted very poorly in bulk soil and was not detected in samples collected over the winter period following release to March 1994 (Table 8.2). The GMM (1.9 x 102 cfu g-1) was isolated from only one soil sample (October 1994) from all of the bimonthly samples collected from each plot between March 1994 until December 1996 (n= 144).

8.6. Survival of GMM on Over-Wintering Sugar beet Secondary Growth, Resown Crop Plants and Volunteer Weeds No GMMs were detected in samples of over-wintered leaf rosettes and the rhizospheres of the sugar beet plants collected in mid March 1994 (11 months after sowing). When samples of new secondary leaf and rhizosphere growth were collected 384 days after sowing of the inoculated seeds, GMMs were detected (Table 8.2). These old plants were removed to prevent bolting and new sugar beet seed was sown in the recombinant treated plots. The new seedlings and subsequent plants were assessed for the presence of the GMM at approximately bimonthly intervals. In spring 1995 the plants were again removed and the plots resown with sugar beet. Subsequent detection of the GMM was transient and highly variable. GMMs were recorded on the secondary leaf growth of the 1993 plants in May 1994 (day 384). However, even in the emerging leaves of an old plant the level of colonization was low, with an average of 1.8 x 103 cfu g-1 in three replicates. This was at least one order of magnitude below the density recorded from plants that had been colonized from inocula introduced with the seed at the time of planting. On the same occasion, the GMM was also detected in rhizosphere soil in one of three replicates at a level of 5.4 x 102 cfu g-1. In June 1994, GMMs were detected at a density of 3.5 x 103 cfu g-1 in only one of nine samples of buttercup taken (Ranunculus spp.), but when buttercup was sampled again in August no GMMs could be isolated. In August and October 1994, the GMM was only isolated from habitats below ground level. In August 1994, the GMM was only found in one of nine sugar beet rhizosphere soil samples at 1.1 x 102 cfu g-1. In October 1994, the GMM was only detected in one of nine sugar beet rhizoplane samples at 8.6 x 101 cfu g-1. The source of the inoculum was unresolved, but the data indicate that the GMM may persist in the phytosphere below the limit of detection (20 cfu g-1) and is able to colonize new plant tissue. No GMMs were isolated from any habitat sampled in December 1994, nor in any sample

Leaf Root Cortex Rhizoplane Rhizosphere soil Secondary leaf Immature leaf Mature leaf Senescent leaf Rhizoplane Rhizosphere soil Leaf Root Bulk soil

Old sugar beet over-wintered from 1993

* * * * * * * * *

March ‘94 +b ++c * * * * * * * *

May ‘94 * * * * * * * * + *

June ‘94 * * * * * * + -

August ‘94 * * * * * + +

October ‘94 * * * * * -

December ‘94

* * * * * -

March ‘95

isolated from any sugar beet planted in April 1995, or from weed or bulk soil sample collected April 1995 - December 1996. The bacterium, P. fluorescens SBW25 was isolated from the leaf surface of sugar beet grown at the field site, Wytham UK, and by site directed homologous recombination was chromosomally marked, one locus carrying lacZY (lactose utilisation and ability to turn X-gal blue) and at another locus with aph-xylE (kanamycin resistance and the ability to turn catechol yellow). In combination these novel phenotypes provided unique and highly sensitive isolation and molecular methods for strain and marker gene detection.

Except for weeds, triplicate samples were taken. Limits of detection 5 cfu g-1 aWeeds: Creeping Buttercup (Ranunculus repens L.), Fat hen (Chenopodium album L.,) Knotgrass (Polygonum aviculare L.) and Thistle (Carduus tenuiflorus Curt.). - No GMM detected in any replicate. *Not determined. bGMM detected in one replicate. cGMM detected in more than one replicate. GMM was not

Weeds a

New sugar beet planted April 1994

Habitat

Plant

Table 8.2. Detection of a GMM, Pseudomonas fluorescens SBW25EeZY6KX, in the sugar beet release plots Wytham, 1994. Inocula applied once as a seed dressing April 1993.

Field Released Pseudomonas fluorescens SBW25EeZY6KX 121

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taken from replanted sugar beet leaves or roots, volunteer weeds or soil samples collected in 1995 or 1996 (Table 8.2).

8.7. Would Persistence of a Modified Indigenous Strain in the Soil Environment Pose a Threat? One of the major concerns about the release of GMMs is their potential to persist and proliferate for long periods of time in the environment. The longer the organism is present in the environment the greater its potential to cause an impact. Therefore, it is essential to assess the longevity of any released organism in order that the risks associated with it can be properly estimated. Sensitive culture based assays were developed which were able to detect 1 cfu g-1 of inocula17 and this approach was preferred to molecular based techniques, including PCR assays of similar sensitivity,24 because of their ease of use in the analysis of the large numbers of samples collected to provide statistically relevant data. In addition, we wanted to assess whether dormant populations or populations persisting at densities below the level of detection could be selected for by the introduction of growing plants. The introduction of plants was used as an environmental enrichment protocol to measure the persistence of viable bacteria rather than the persistence of nucleic acids. In the field investigations undertaken in 1993, the inocula persisted at low levels in the soil and was able to colonize the sugar beet plants during the growing season. At the time of harvest, associated with the first hard frosts, the population of the introduced GMM had declined considerably.15 However, in the following spring of 1994 secondary plant growth from over-wintering sugar beet apparently supported a transient GMM population which did not establish a sustained GMM community and was not found on mature or immature leaves sampled 60 days later. Further evidence that the GMM was dependent on plant material for over-winter survival, was shown as the GMM colonized wheat roots during 1993-1994 at densities as high as 106 cfu g-1 roots, but could not be isolated once roots could no longer be recovered from the soil seven months after harvest.21 The limited amount of dissemination and persistence of P. fluorescens SBW25EeZY6KX (the GMM) in soil is in agreement with other similar studies.19,25 These investigators monitored the survival of a P. fluorescens, marked with lacZY, in the rhizosphere of wheat and concluded that there was limited lateral movement and a failure of inocula to persist or colonize subsequent introduced crops. If precautions are necessary, then the removal of plant material from release sites for at least one season postrelease might be appropriate. This assumption would need to be tested. Other bacteria better adapted to harsh soil conditions may survive more effectively in bulk soils. For example, rhizobia have persisted at ecologically significant levels at Rothamstead Research Station (UK) since their release in 1987.26 The highly specialized nature of these nodulating bacteria as well as their ability to survive in soil no doubt has selected for strains that are able to persist to provide inocula able to colonize appropriate host plants as they develop. Although no genetic instability or mobilization of marker genes from the GMM could be demonstrated in vitro or as a result of the field releases,14,15,18 it was established that the colonizing populations of the GMM interacted with the indigenous microflora in the exchange of genetic material in vivo4 and that carriage of novel plasmids affected the ecology of the bacterial host.5 Therefore, it was important to establish whether any isolates detected during the postrelease monitoring of soils and volunteer plants had acquired these or other plasmids, or lost or transferred their markers to other bacteria. In all samples collected, no evidence for plasmid transfer to the GMM or persistence of GMM transformants was gathered. All isolates retained the complete GMM phenotype, i.e., they produced blue colonies on media supplemented with kanamycin and X-gal (genotype; lacZY, aphI, xylE). No kanamycin resistant, white colonies (lacZ-) were identified that reacted with catechol. Such reac-


123

tions would have identified mutants or indicated the selection of transconjugants as the kanamycin resistance-catechol 2,3 dioxygenase genes were linked and inserted at the same chromosomal site in P. fluorescens SBW25EeZY6KX. Small populations of the GMM probably survived under particular conditions as colony forming units were detected at highly variable densities on a few samples collected from new plant growth and rhizosphere soil samples. However these populations were transient and failed to become established. The loss of the inocula in a viable state from the release site was confirmed in all subsequent surveys conducted in 1995 and 1996. The absence of the introduced bacteria, as determined by plate counts assay and PCR screening of total extracted soil DNA,14,24 or any evidence for gene transfer, confirms a lack of sustained activity in the environment. In the absence of activity it can be assumed that negligible or no risk occurred to the environment as a result of these investigations. So what are the risks associated with GMMs entering the environment? Should they be considered more or less hazardous than the current usage of chemical and biological reagents? To make such assessments, we must provide data drawn from a sound basic scientific understanding, which addresses concerns, real or potential, from which accurate predictions can be made in order to gain public confidence and (if possible) identify and quantify hazards.

8.8. Conclusion Irrespective of the origin of the inoculum, the introduction of a large number of organisms to any habitat is likely to have some effect (impact), even if only short term. This ‘inoculum effect’ has been demonstrated with both the introduction of unmodified bacteria and GMM’s. Unmodified microbes have been released into the environment for many decades, particularly in the agricultural practices of the biocontrol of pathogens and inoculation of crop plants with nitrogen fixing micro-organisms. Few studies have attempted to determine the effect of these introductions on indigenous microbial communities. Assessments of the risks from GMMs have been made in situ in field releases where inocula have been released into the natural environment.13,15,25,28,39,30 Particular consideration has been given to the potential impact GMM’s may have on ecosystems by disturbing natural communities or by exchanging genetic material.28,31 The plant surface and rhizosphere are the target of a number of releases and have been the focus of many studies, including those of fluorescent pseudomonads in the rhizosphere of a number of crops.15,18,19,21,25,30,32 Typically, GMMs contain introduced antibiotic resistance genes which facilitate their detection against the background of indigenous strains sampled form the environment, and as a consequence of the technical approaches used to generate the recombinants. It should be emphasized that the use of antibiotic resistance genes in GMMs does not in itself pose an unacceptable generic risk. Potential harm may only arise from the transfer of the introduced marker genes to other bacterial populations where the incidence or likelihood of external selective pressures (e.g., antibiotics) are encountered by the GMMs and indigenous suitable recipient populations. Obviously, the use of marker genes that impart resistance to clinically important antibiotics would not be advised, but the use of a marker resistance in a GMM, for example kanamycin or ampicillin, where the specific phenotype is comparatively common in soils and other natural environments, would not be considered as an additional serious hazard. The possibility of selection through the use of antibiotics in animal health or growth promotion could be a factor (Chapter 2). This may be compounded by the application of untreated sewage or slurry to land where the GMM has been applied or might be encountered. As with current practices, all GMM releases should be taken on a case-by-case basis to evaluate their consequences on entering the environment, a part of which is the consideration of the construct and the nature of the introduced genes themselves.

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The nature of the introduced genes and the resulting phenotype is relevant as this will influence the degree of risk. The advantage of selective markers is that they can be used to readily track and monitor the fate, persistence and activity of released GMMs. The facility to track inocula, and therefore the fate of the genes is an important safety consideration. Manipulated genes can be transferred horizontally to or from a GMM where they may persist and be expressed. Although the frequencies of horizontal transfer will usually be low, the location of manipulated genes at carefully chosen chromosomal sites can considerably reduce these events further. However, it should not be necessary to prescribe the chromosomal location of introduced genes unless minimizing the chance of gene transfer is preferred. Investigations with marked mobile elements will provide data to evaluate gene flow in the environment to ascertain whether restrictions to the chromosome, avoiding all known mobile genetic elements is appropriate. The nature of the construct will dictate the probability of gene transfer, but perhaps the primary consideration must remain the phenotype of the introduced genes and how the ecology and activity of the modified host is altered. However, any introduced genes, in order to persist and proliferate, require at the very least periodic amplification (selection) of their host. With the decline in concentration of the manipulated genes in the environment, the potential for the future proliferation and spread of these genes is considerably reduced. Precise prediction of the changes in behavior arising from the acquisition of new genes is especially difficult because microbial populations are engaged in highly complex interactions. Assessment of risks arising from the transfer of manipulated genes requires the nature and activity of the genes selected for use in a GMM to be considered on a case by case basis. From the results presented in this chapter it would appear that the persistence of the released GMM, P. fluorescens SBW25EeZY6KX, in the environment is limited. Even when the GMM was detected it was present at very low levels and not in all replicate samples. This lack of consistency may reflect that the levels of GMM were close to the limits of detection. Lateral dispersal of the GMM during the release was limited so it was probable that the GMM was confined to a relatively small area of soil around the plant root. Given this scenario, it was unlikely that colonization of plant roots would occur unless seeds were sown in the exact position where an inoculated plant had grown the previous season. Thus, the numbers of GMMs detected in 1994/5 were not considered to be ecologically significant. This view was supported by our inability to detect the GMM in 1996. This implies that, as P. fluorescens SBW25EeZY6KX does not appear to be able to persist in soil, it will not have any long-term effects upon the site to which it is released.

Acknowledgments These investigations were supported by the UK Department of the Environment and the EU-Biotech program. Mark J. Bailey is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Ellis RJ, Thompson IP, Bailey MJ. Temporal fluctuations in the genotypic composition of a population of leaf-associated pseudomonads. FEMS Microbiol Ecol 1999; 28:345-356. 2. Thompson IP, Bailey MJ, Fenlon JS et al. Quantitative and qualitative seasonal changes in the microbial community from the phyllosphere of sugar beet (Beta vulgaris). Plant and Soil 1993; 150:177-191. 3. Rainey PB, Bailey MJ, Thompson IP. Phenotypic and genotypic diversity of fluorescent pseudomonads isolated from field grown sugar beet. Microbiol 1994; 140:2315-2331. 4. Lilley AK, Bailey MJ. The acquisition of indigenous plasmids by a genetically marked pseudomonad population colonizing the phytosphere of sugar beet is related to local environmental conditions. Appl Environ Microbiol 1997; 63:1577-1583.


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5. Lilley AK, Bailey MJ. Impact of pQBR103 acquisition and carriage on the phytosphere fitness of Pseudomonas fluorescens SBW25: burden and benefit. Appl Environ Microbiol 1997; 63:1584-1587. 6. Lilley AK, Young JP, Bailey MJ. Bacterial population dynamics: plasmids a genetic mechanism for maintaining diversity and adaptation. In: Thomas CM ed. Chapman and Hall. 1999:In Press. 7. Lenski RE. Quantifying fitness and gene stability on micro-organisms. In: Ginzburg LR. Ed. Assessing risks of biotechnology. Butterworth-Heineman, Boston. 1991:173-192. 8. Ellis RJ, Thompson IP, Bailey MJ. Metabolic profiling as a means of characterising plantassociated microbial communities. FEMS Microbiol Ecol 1995;16:9-18. 9. De Leij FAAM, Sutton EJ, Whipps JM et al. Effect of a genetically modified Pseudomonas aureofaciens on indigenous microbial populations of wheat. FEMS Microbiol Ecol 1994; 13:249-258. 10. Doyle JD, Stotzky G. Methods for detection of changes in the microbial ecology of soil caused by the introduction of micro-organisms. Microbial Releases 1993; 2:63-72. 11. Weller DM. Colonisation of wheat root by fluorescent pseudomonads suppressive to takeall. Phytopathol 1993; 73:1548-1553. 12. Yuen GY, Schroth MN. Interactions of Pseudomonas fluorescens strain E6 with ornamental plants and its effect on the composition of root colonizing microflora. Phytopath 1986; 76:176-180. 13. Thompson IP, Ellis RJ, Bailey MJ. Autecology of a genetically modified fluorescent pseudomonad on sugar beet. FEMS Microbiol Ecol 1995; 17:1-14. 14. Bailey MJ, Lilley AK, Thompson IP et al. Site directed chromosomal marking of a fluorescent pseudomonad isolated from the phytosphere of sugar beet; stability and potential for marker gene transfer. Molec Ecol 1995; 4:755-764. 15. Thompson IP, Lilley AK, Ellis RJ et al. Survival, colonisation and dispersal of genetically modified Pseudomonas fluorescens SBW25 in the phytosphere of field grown sugar beet. Bio/Technology. 1995; 13:1493-1497. 16. Rainey PB, Bailey MJ. Physical and genetic map of the Pseudomonas fluorescens SBW25 chromosome. Mol Microbiol 1996; 19:521-533. 17. De Leij FAAM, Bailey MJ, Lynch JM et al. A simple most probable number technique for the sensitive recovery of a genetically engineered Pseudomonas aureofaciens from soil. Lett Appl Microbiol 1993; 16:307-310. 18. Bailey MJ, Lilley AK, Ellis RJ et al. Microbial ecology, inocula distribution and gene flux within populations of bacteria colonizing the surface of plants: Case study of a GMM field release in the UK. In: Van Elsas JD, Trevors JT, Wellington EMM eds. Modern Soil Microbiology. Marcel Dekker, New York. 1997:479-500. 19. Drahos DJ, Barry GF, Hemming BC et al. Spread and survival of genetically marked bacteria in soil. In: Fry JC, Day MJ, eds. Release of genetically engineered and other microorganisms. Cambridge University Press, Cambridge. 1992:147-159. 20. De Leij FAAM, Sutton EJ, Whipps JM et al. Spread and survival of a genetically modified Pseudomonas aureofaciens on the phytosphere of wheat and soil. Appl Soil Ecol 1994; 1:207-218. 21. De Leij FAAM, Sutton EJ, Whipps JM et al. Field release of a genetically modified Pseudomonas fluorescens on wheat: Establishment, survival and dissemination. Bio/technology 1995; 13:1488-1992. 22. Lilley AK, Hails RS, Cory JS et al. The dispersal and establishment of pseudomonad populations in the phyllosphere of sugar beet by phytophagous caterpillars. FEMS Microbiol Ecol 1997; 24:151-158. 23. Bailey MJ, Thompson IP. Detection systems for phylloplane pseudomonads. In: Wellington EMH, van Elsas JD, eds. Genetic interactions between micro-organisms in the environment. Pergamon Press 1992:126-141. 24. Bramwell PA, Barallon RV, Rogers HJ et al. Extraction and PCR amplification of DNA from the rhizoplane. In: Akkermans ADL, Van Elsas JD, DeBruijn FJ, eds. Molecular Microbial Ecology Manual. Kluwer Academic Publishers. 1995:36-55.

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25. Kluepfel DA, Kline EL, Skipper HD et al. The release and tracking of genetically engineered bacteria in the environment. Phytopathol 1991; 81:348-352. 26. Hirsch P, Spokes JD. Survival and dispersion of genetically modified rhizobia in the field and genetic interactions with native strains. FEMS Microbiol Ecol 1994; 15:147-160. 27. Denning N, Morgan J, Whipps JM et al. The flagellin gene as a stable marker for detection of Pseudomonas fluorescens SBW25. Lett Appl Microbiol 1997; 24:198-202. 28. Bailey MJ, Lilley AK, Thompson IP et al. Deliberate release of Recombinant Micro-organisms. In: Demain AL, Davies JE, eds. Manual of Industrial Microbiology and Biotechnology 2nd edition. American Society Microbiology, Washington. 1999:693-703. 29. Cory JS, Hirst ML, Williams T et al. Field trial of a genetically improved baculovirus insecticide. Nature 1994; 370:138-140. 30. Wilson M, Lindow SE. Release of recombinant micro-organisms. Ann Rev Microbiol 1993; 47:913-944. 31. Tiedje JM, Colwell RK, Grossman YL et al. The planned introduction of genetically engineered organisms—Ecological considerations and recommendations. Ecology 1989; 70:298-315. 32. De Leij FAAM, Sutton EJ, Whipps JM et al. Field release of genetically modified Pseudomonas fluorescens on wheat: Impact on indigenous microbial populations. Appl Environ Microbiol 1995; 61:3443-3453.

CHAPTER 9

Use of luc-Tagged Genetically Modified Microorganisms (GMMs) to Study Rhizobial Ecology in Soil Columns, Field Lysimeters and Field Plots Christoph C. Tebbe

9.1. Introduction

T

he benefit and importance of genetically tagged microorganisms for agricultural biotechnology can be demonstrated with studies on plant-microbe interactions such as the symbiosis between rhizobia and leguminous plants (see also Chapter 11). Rhizobia, a group consisting of the five genera Rhizobium, Sinorhizobium, Mesorhizobium, Azorhizobium, and Bradyrhizobium1 are soil bacteria which are able to infect roots of specific leguminous host plants and induce the formation of root nodules into which the transformation of atmospheric nitrogen (N2) into organic nitrogen takes place (nitrogen fixation). Rhizobia provide the essential enzymatic machinery for this process, most importantly the nitrogenase enzyme, whereas the plant supplies the intracellular bacteria (bacteroids) with photosynthetically generated organic nutrients. Additionally, the plant tissue protects the bacteroids against oxygen, a compound that inhibits the nitrogenase. By this symbiosis, plants become independent of mineral or organically dissolved nitrogen sources. On the other hand, the symbiosis is inhibited if sufficient bound nitrogen is available for plant growth. In agriculture, nitrogen fixing crop plants, such as alfalfa, do not require fertilization with organic or mineral nitrogen. Such plants can be included into crop rotations to increase the nitrogen content of soil and thus optimize conditions for the following plant, e.g., wheat. However, if no symbiotic partner bacteria or bacteria with low nodulation efficiencies are present in the soil, leguminous plants may fail to fix nitrogen. To overcome this risk, leguminous plants have been inoculated with rhizobia in agriculture for more than 100 years. However, in some cases bacterial inocula may fail to induce root nodule formation and nitrogen fixation.2 One reason is that the commercially available inoculants may have a poor quality due to the presence of contaminant, nonrhizobial, bacteria.3 But even if pure cultures of potentially effectively nodulating and nitrogen-fixing bacteria are utilized in field applications, beneficial effects such as shown by enhanced growth of leguminous host plants are not guaranteed. Nonefficient inoculant preparation or application may be Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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one reason for the lack of success,4,5 but the main reason seems to be the unpredictable impact of environmental factors (e.g., temperature, humidity, nutrient availability) upon the released bacterial cells, or the presence of other plant roots or indigenous competing strains of the same species as the inoculant.6 In most cases of unsuccessful field applications, it is not known whether the lack of plant growth promotion can be attributed to the poor survival of the inoculant, the loss of its competitive capability for nodulation, or to other factors.7 To evaluate the reason for such failures it would be helpful to analyze the rates of survival of such inoculants. Although several methods for reisolation, identification and quantification of rhizobia from soil are available, most of them require extensive laboratory studies, e.g., nodulation of host plants in the laboratory or greenhouse followed by genetic fingerprinting, to distinguish inoculated cells from indigenous cells of the same species.8-10 Due to the methodological efforts, the number of samples which can be analyzed in one laboratory is rather small and may not be sufficient for the evaluation of a specific inoculant in field release experiments. Marker gene technology can provide the ideal tools to perform such evaluation studies, since an inoculant can be tagged specifically with a quickly detectable and unambiguous trait.11,12 The characteristics, potentials and limitations of these tools are described in detail in Chapters 5-7. Depending on the marker gene, strains can be directly detected by microscopy (without cultivation), by PCR amplification of such genes from DNA directly extracted from microbial communities (see also Chapter 3), or by quantification after growth (colony formation) on selective or nonselective growth media (Chapter 2). An important point to consider before using marker gene -tagged strains for evaluation as inoculants is, that the insertion of a marker gene or its expression may affect the fitness of a strain and thereby decrease the capacity to compete with indigenous bacteria.13-15 Another potential disadvantage for field applications of marker gene-tagged strains is, that they are “genetically engineered” (genetically modified microorganisms; GMMs) and, thus, their release requires thorough risk assessment studies and permission of competent national authorities. In fact, the case study reported in this chapter included the first two field releases of GMMs in Germany. These releases were conducted in a collaboration between the University of Bielefeld (M. Keller, A. Pühler) and the FAL (Forschungsansalt fuer Landwirtschaft; C.C. Tebbe, J.C. Munch).

9.2. Construction and Properties of GMM Strains L1 and L33 In order to perform a field study with Sinorhizobium meliloti (formerly: Rhizobium meliloti), two different marker gene-tagged strains were constructed at the University of Bielefeld, Germany. S. meliloti nodulates the pasture crop alfalfa (Medicago sativa) and also weed plants of the genus Melilotus and Trigonella. The nonengineered strain S. meliloti 2011 was streptomycin resistant (intrinsic marker). By cultivation of soil suspensions on selective, streptomycin amended growth media (agar plates; “selective plating”), the background population of co-cultivated soil microorganisms could be reduced approx. 100-fold and thus, the sensitivity of GMM detection was enhanced.16 The marker gene used for construction of the engineered strains in this study was the luciferase gene (luc). This gene was selected, since the metabolic burden imposed by this gene upon the organism was expected to be low and thus, the fitness of tagged strains would not be affected compared to that of the wild-type strain (see also Chapter 5). The luc reporter gene was combined with a constitutively expressed broad host range promoter (nptII), isolated from transposon Tn5. This promoter was selected to allow efficient detection of marker gene expression and potentially to also detect the marker gene in other soil bacteria, as a result of intergeneric gene transfer. The marker gene was detected after blotting colonies grown on plates onto nylon membranes, wetting these membranes with a luciferin solution and detecting light emis-

Use of luc-Tagged GMMs to Study Rhizobial Ecology

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sion in a film cassette. It is important to notice, that, due to its oxygen demand, luciferase gene expression could only be detected when air was permitted to diffuse between the attached colonies and the film material. This was achieved by inserting a distance holder of 0.5 cm into the film cassettes. In strain S. meliloti L1, the combined nptII-luc gene construct was chromosomally inserted into the recA gene. This gene is involved in homologous recombination of DNA and also in DNA repair mechanisms, such as induced after UV exposure.17 Since the luc gene cassette was inserted into this gene, the gene sequence was disrupted and, therefore, strain L1 exhibited a RecA- phenotype.16 Microcosm studies in the laboratory demonstrated that in the absence of its host plant, alfalfa, the recA-mutation reduced the ability of strain L1 to persist in bulk soil.18 The other strain selected for our field release study was an isogenic strain of S. meliloti 2011. This strain, designated L33, contained the same reporter gene cassette as S. meliloti L1, but its chromosomal insertion was downstream of the recA gene, and, thus, not disrupting it (Fig. 9.1). The strain was indistinguishable from its parent strain 2011, except for the expression of the marker gene.19

9.3. Experimental Design: What Can Be Achieved with Each System? Three different successive experimental setups were used at the FAL (Braunschweig) to study the survival and ecological fitness of tagged strains, S. meliloti L1 and L33 (Fig. 9.2). The first experimental setup consisted of soil columns in the greenhouse, the second were field lysimeters of the same size as the greenhouse soil columns and the third setup were field plots which were inoculated with the respective S. meliloti strains. Characteristics of the three setups are shown in Table 9.1. The experiments were started in three successive years (1993 to 1995).

Fig. 9.1. Genetic maps of the chromosomal recA regions in the S. meliloti (former name: Rhizobium meliloti) strains 2011 (wild-type), L33 and L1 and positions of primers, which allow to differentiate L1 and L33 by size of the PCR product. Reprinted with permission from: DammannKalinowski T, Niemann S, Keller M et al. Appl Microbiol Biotechnol 1996; 45:509-512.

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Table 9.1. Comparison of microcosms, lysimeters and field plots used to study the survival and ecological interactions of Sinorhizobium meliloti strains in soil Parameter

Greenhouse soil columns

Field lysimeters

Field plots

Size

diameter: 30 cm depth: 65 cm

diameter: 30 cm depth: 65 cm

3 m x 3 m squares no depth limitation

soil horizons

reconstructed, equilibrated for 2 month

reconstructed, equilibrated for 1 year

natural horizons (plough layer 25 cm depth)

number of 3 replicates for each treatment

4

5

total surface 0.4 m2 area inoculated with each strain

0.6 m2

45 m2

total number approx. 3 x 1011 of genetically engineered cells released

approx. 1 x 1011

approx. 2 x 1013

inoculation technique

mixing of bacterial cell suspension and peat into the upper 4 cm of the soil

mixing of bacterial cell suspension into the upper 4 cm of soil in the laboratory and subsequent transfer onto field lysimeters

spraying of cell suspension onto the soil surface

Monitoring parameters

* survival of GMMs * recombinant gene persistence * nodulation efficiency * GMMs colonization of different soil horizons * effect on plant growth * effect on organic carbon and nitrogen concentrations in soil * microbial biomass * quantification of selected culturable bacterial populations * immediate metabolic response (“Biolog-method”)

* survival of GMMs * * nodulation efficiency * * GMMs colonization of different soil horizons * vertical transport of * GMMs * effect on plant growth * effect on organic car- * bon and nitrogen concentrations in soil * * microbial biomass * quantification of selected culturable bacterial populations *

survival of GMMs; horizontal transport of GMMs rood nodule occupancy in the field nodulation efficiency colonization of rhizospheres from host plants and weed impact of GMMs and the microbial diversity found in plant rhizospheres * ingestion and transport of GMMs by soil invertebrates


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Parameter

Greenhouse soil columns

Field lysimeters

Field plots

stability of ecological parameters

Low

Medium

High

maximum length of monitoring

1.5 years

2 years

> 3 years

Problems

* soil compaction * crevices in soil columns and rim effects * limited amount of sampling dates

* limited amount of sampling dates * refilling holes after auger insertions and its impact on soil structure

* large scale production of inoculants * inoculation of field plots without aerial spread of GMMs * spread into neighboring plots with host plants

Reference

Schwieger et al23

Schwieger et al24

Dresing et al25 Schwieger and Tebbe26

The major difference between soil columns in the greenhouse and field lysimeters were the environmental factors acting on both systems. In the greenhouse, soil columns were wetted to allow plant growth. However, in order to study active movement of the GMM strains, columns were not saturated with water and, thus, no flow-through water could be collected. In order to protect the columns from frost damage, the minimum temperature in the greenhouse was not permitted to be below 4°C. Moreover, the maximum temperature was not above 30°C. In contrast, field lysimeters were exposed to the natural conditions, including periods with temperatures below 0°C and above 30°C. Wind exposure in the field resulted in a rather quick drying of the soil in the lysimeters, even after heavy rain periods. On the other hand, large amounts of flow-through rain water could be collected; 42.5 l per lysimeter over a period of 18 months. The soil surface inoculated in the field plot investigation was approx. 100 times larger than that of the greenhouse columns or field lysimeters. This allowed the removal of more material for analyses during this investigation. Also, holes created by auger insertions in the plot experiment during sampling were not refilled, as in greenhouse or lysimeter studies, since vertical transport was not analyzed and such holes were not much different in size compared to those, naturally produced by mice in the field. The selection of parameters monitored along with the assessment of the survival of the inoculated S. meliloti strains L1 and L33, respectively, depended on the characteristic specificities of each model system. Since the model systems were studied subsequently, we could optimize our monitoring methods and omit parameters that proved to be insensitive or remained unaffected. For instance, the immediate metabolic response of soil extracted microbial communities20,21 (community level physiological profiles, CLPP) did not detect any differences between the treatments (inoculation, noninoculated controls). Therefore, this parameter was not further included in the field monitoring. Transport by rain water could not be analyzed on the field plots, but on the other hand, the plots were ideal for study of the colonization of rhizospheres of host plants and weed plants growing between the host plants. Also, only from the field plots, a large variety of soil insects could be collected

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during the growing season. The gut contents of several of these insects were analyzed for the occurrence of marker gene tagged, bioluminescent cells.22

9.4. Techniques and Consequences of GMM Soil Inoculations In all three experimental setups it was intended to start the experiment with approx. 106 cells of the strain g-1 soil in the upper soil horizon (plough layer; 0 to 25 cm depth). In greenhouse experiments, batch culture grown cells were harvested, washed in potassium phosphate buffer (50 mM, pH 7.2), and mixed with sterile peat and soil taken from the upper 4 cm of the soil columns. This mixture was loaded onto the soil columns and alfalfa seeds were added. A similar technique was applied to inoculate the field lysimeters. However, we abstained from utilizing peat as the carrier, since laboratory studies did not show any beneficial effect of peat on the establishment of S. meliloti in the field soil. Also, to study the survival and vertical transport of inoculated cells we decided to exclude peat as a material, since peat would have provided additional nutrients and would have protected the inoculated cells against predation. Finally, vertical transport was a monitoring parameter in this study and adsorption of the inoculated cells to the peat would have prevented translocation of such cells into deeper soil horizons. Due to the size of the field plots that had to be inoculated, the surface soil could not be removed and inoculated in the laboratory. Instead, a spraying machine, normally used to apply pesticides onto experimental field plots, was utilized and bacterial suspensions were directly sprayed onto the soil surface in the field. In contrast to the other systems, aerial spread of the inoculant into neighboring field plots could not be excluded even though the spraying machine was modified by utilizing a rather low pressure (1 bar). Also, a specifically developed box only allowed bacterial cells to move downwards but not sidewards (see Fig. 9.2, right side). During the inoculation period the wind strength was low (0.5-3 ms-1). The concentration of inoculated cells (106 cfu g-1 in the plough horizon) was one to three orders of magnitude above the upper limit of rhizobial populations found in well colonized soils.6,7 Compared to the total number of bacterial cells commonly found in soil, the inoculant was less than 1% of the population. With the three experimental settings used in this study (soil columns, field lysimeters, field plots), the titers of inoculated cells, as determined immediately after the beginning of each experiment varied by one order of magnitude (Table 9.1). However, as shown by the survival of both strains, S. meliloti L1 and L33, the fate of the release strain was not dramatically influenced by the number of

Fig. 9.2. Three model systems used to study the survival and microbial ecology of bioluminescent S. meliloti strains (GMMs) in soil. Left, soil columns in the greenhouse, allowing to study bacterial colonization of different soil horizons; field lysimeters (middle) of the same design, and field plot inoculation (right).


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cells initially introduced into each model system. Due to the relatively low amount of microbial cells added, it was not surprising to see that general parameters, like microbial biomass, organic carbon or total nitrogen (with variances of at least ± 10%), did not respond to this treatment at all. Other parameters, like population sizes of culturable heterotrophic bacterial communities on cellulose, glucose or aromatic compounds, also did not respond to the inoculation procedure. Thus, these parameters, which were suspected to be indicative for nonintended, dramatic changes within the soil microbial community, were only monitored during the first two stages of this investigation (soil columns and field lysimeters).

9.5. Survival and Spread of GMMs in Soil columns, Field lysimeters and Field Plots In the greenhouse experiment, a continuous decline of the inoculated cells was observed over a period of 85 weeks (Fig. 9.3). Strain S. meliloti L33 declined to 9.0 x 104 cfu g-1 soil within 24 weeks and to 2.8 x 103 cfu g-1 within 85 weeks in the upper 25 cm of the soil columns.23 Decline rates for S. meliloti L1 were not significantly different, indicating that in the presence of its host plant, alfalfa, and under greenhouse conditions, the recA-mutation did not affect the environmental fitness of S. meliloti. In order to confirm that the decrease of the GMM was due to its elimination from the soil habitat, and not just a decrease of culturable cells (for more details of this phenomenon, see Chapter 1), we additionally monitored the presence of the luc marker gene in DNA directly extracted from soil by PCR. Techniques to obtain DNA from soil suitable for PCR detection are described in Chapter 3. Using carefully selected primers, we could differentiate between product sizes obtained from strain L1 and L33 (for positions of primers, see Fig. 9.1). By this means we were able to show that no cross-contamination of inoculated GMM cells occurred between the lysimeters (Fig. 9.4). Additionally, following GMM inoculation, a decrease in PCR product yields was observed. This decrease was probably caused by a decline of the template copy numbers in soil DNA. A similar decrease of inoculated cells was detected after field lysimeter inoculation: after 24 weeks, 2.0 x 104 cfu g-1 soil were detected for strains L1 and L33, respectively. In contrast to the greenhouse experiment, however, the titer did not decrease any further, until growth of alfalfa was stopped after 80 weeks.24 A similar population decline as observed in the field lysimeters, was detected after inoculation of field plots. Populations of both strains dropped from 106 to below 104 within 14 weeks, then increased to 2 x 104 cfu g-1, 24 weeks after inoculation. Until today, four years after the field release, the titers of L33 and L1 Fig. 9.3. Survival of S. meliloti strains in the upper 25 cm in soil columns (greenhouse). Strain L1 (▲) and strain L33 (●). Reprinted with permission from: Schwieger F, Willke B, Munch JC et al. Biol Fertil Soils 1997; 25:340-348.

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Fig. 9.4. PCR mediated detection of the recombinant luc-marker gene in DNA, directly extracted from soil. PCR products of strain L33 (415 base pairs; lanes 3, 5, 7, 9, 11, 13, 16) and L1 (1011 base pairs; lanes 2, 4, 6, 8, 10, 12 and 15) are clearly distinguishable. DNA was extracted from soil columns (0-25 cm depth) after a day (lanes 2, 3), 2.1 weeks (4, 5), 4.1 weeks (6, 7), 8.1 weeks (8, 9), 16.1 weeks (10, 11), and 24.1 weeks (12,13). Other lanes show size standards and controls. Reprinted with permission from: Schwieger F, Willke B, Munch JC et al. Biol Fertil Soils 1997; 25:340-348.

remained in the range between 2 x 103 and 7 x 104 cfu g-1 soil, with a seasonal impact on the population sizes. Interestingly, in the first two years, the populations of L1 were below those of L33 in Fall and Winter but above L33 in Spring, as observed in the two following years.25 This suggested that there was an ecological significance of the recA mutation. However, in the third year, this phenomenon was not significant. Even though it is extremely difficult to determine the reason for significant differences between recA- and recA+, the majority of sampling dates did not yield such differences and, thus, we can conclude that the recA gene was not crucial for S. meliloti to successfully colonize the field plots with alfalfa. Due to the high persistence of both strains, L1 and L33, we can also conclude that the luciferase marker gene did not interfere with the environmental fitness of S. meliloti. This clearly supports the assumption that in contrast to some other marker genes, such as luxCDABE, luc has a rather low impact on fitness. Vertical transport of surface soil inoculated cells was studied in greenhouse columns and field lysimeters, but only the latter system yielded reliable data. In the greenhouse, more than 98% of the inoculated cells were recovered in the upper 10 cm in three of four soil columns analyzed after 85 weeks of incubation.23 However, in one column, layers below 20 cm soil depth were almost homogeneously colonized with titers of 104 to 105 cfu g-1 soil. Rim effects and crevices in the soil column probably led to transport of the surface inoculated cells in that column. Additionally, oxygen diffusion from the bottom of the soil columns, which could not completely be sealed from the greenhouse atmosphere and a homogenous temperature of the soil column, may have promoted growth in such “deeper” soil layers. In the lysimeters, the temperature and the gas atmosphere were more similar to conditions in the surrounding field soil environment. With this, more realistic system, monitoring of the inoculated cells showed no migration of the GMMs into layers below 20 cm depth (threshold of detection 100 cfu g-1 soil) and flow-through rain water did not transport any detectable bioluminescent cells through the 65 cm soil profile (threshold of detection 10 cfu ml-1). Thus, it could be concluded that no risk of vertical migration of GMMs on the field site for the subsequent field plot inoculation existed. By selection of the field site, at


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which the groundwater table was 20 m below the soil surface, unintentional spread into ground water could be excluded. Horizontal spread of inoculated cells could only be analyzed in the field plot experiment. The experimental field consisted of 20 plots, each a square of 9 m2 in 4 x 5 rows, with each plot separated from the other by 3-meter noninoculated strips, seeded with grass. Plots were inoculated in block randomized order with S. meliloti wild-type, L33, L1, or not inoculated. Already 12 weeks after the field release, bioluminescent cells were detected in the rhizosphere of alfalfa growing on noninoculated plots.26 Two weeks later, when we analyzed the titer of bioluminescent cells in bulk soil, recombinant cells were detected on the noninoculated control plots with an average titer of 2.2 x 101 cfu g-1 soil. This titer increased further throughout the 3 year monitoring period to numbers only one order of magnitude below those on the inoculated plots.25 Mixed populations of strain 2011, L33 and L1 were found on wild-type-inoculated plots. Thus, inoculation of S. meliloti 2011 did not completely inhibit the colonization by the GMMs. Sampling outside of the alfalfa seeded plots never resulted in detection of any significant amounts of bioluminescent cells.25 The horizontal spread of the GMMs was obviously restricted to the presence of alfalfa roots. Unintentionally, a large number of different weed plants (approx. 20 different species) grew on the alfalfa-seeded plots during the first vegetation period after inoculation. Several of such weed plants were sampled concomitantly with alfalfa plants to study their rhizosphere colonization by bioluminescent cells. As expected, the rhizosphere of alfalfa plants provided a well suited habitat for S. meliloti and was densely colonized by bioluminescent cells (> 105 cfu g-1 root material). Some weed plants, e.g., Capsella bursa-pastoris, did not enrich for any bioluminescent rhizobial cells, but for the weed Chenopodium album, we found approx. 103 cfu g-1 root material on inoculated and 101 cfu g-1 on noninoculated control plots, 12 weeks after the field release. A total of approx. 1,200 pure culture colonies were isolated on growth media adapted to the isolation of rhizosphere bacteria. These isolates were obtained from rhizospheres of alfalfa and C. album plants, grown on inoculated and noninoculated plots 12 weeks after the release of the GMMs. The species richness, as detected by restriction fragment length polymorphism of PCR amplified 16S rRNA genes (ARDRA) was higher in rhizospheres of alfalfa than of C. album. The diversity of isolates was characterized at the phylogenetic level using restriction fragment length polymorphisms of PCR amplified 16S rRNA genes. The number of ARDRA pattern types, which correlated with species richness and diversity (expressed as the Shannon Index), was larger in rhizospheres of alfalfa than in that of C. album. The species richness was unaffected by inoculation in rhizospheres of C. album but increased in rhizospheres of alfalfa.26 Possibly, the S. meliloti inoculation increased the nutritional status of the early developing alfalfa plants and concomitantly resulted in the release of rhizosphere exudates stimulating the growth of a larger variety of soil bacteria.

9.6. Conclusion The results of this still ongoing investigation on the ecology of a genetically-tagged Sinorhizobium sp. in the field demonstrate the advantages and disadvantages of different model detection systems for such studies. In general, greenhouse soil columns were useful to demonstrate that the model strain, S. meliloti, did not cause any dramatic alterations in the soil microbial community. However, to study vertical translocation and long term effects of inoculated bacterial cells in the soil, more sophisticated systems with temperature gradients, freeze thawing cycles to prevent soil compaction, and control of the soil atmosphere would be needed. Such factors were intrinsically provided in our field lysimeter investigation. The major limitation of both soil columns and lysimeters compared to the field plot experiment was that the number of samples which could be taken for analysis was

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limited due to the small size of both systems. Field plots were also found to be extremely useful for observing the impact of the nondeliberately controlled biological parameters, namely soil invertebrates and weed plants, on the survival of the released GMMs. Interactions with both compartments, which serve as habitats of a large, mainly uncharacterized, diversity of microorganisms, could also be studied. It can be expected, that future environmental applications of recombinant bacterial inoculants will not only include strains for biofertilization like S. meliloti in alfalfa fields, but also bacterial strains for biocontrol or biodegradation. In general, it would be ecologically sound if any GMM construct is eliminated from the environment after its job has been done. A continued persistence potentially includes ecological risks, e.g., exclusion of other indigenous microorganisms, spread into neighboring nontarget ecosystems, transfer of recombinant genes to indigenous microorganisms and noncontrolled establishment of new properties in microbial communities, such as enhanced emergence of resistances. In order to develop GMMs with improved ecological properties, long-term studies on the environmental fate of GMMs are needed. In our own studies, the luc-tagged GMMs were persistent for at least four years after their field release and it can be concluded that this recombinant trait did not dramatically effect the environmental fitness of the strains. Thus, for longterm monitoring of field released microorganisms, the luc marker gene should be an ideal tool.

Acknowledgments I would like to thank Frank Schwieger, Birgit Willke and Rona Miethling for their collaboration which was essential for the success of the project. The co-operation with our colleagues from the University of Bielefeld, especially Mathias Keller, Werner Selbitschka and Alfred Pühler is gratefully acknowledged. Excellent technical assistance was provided by Simone Dose and Phan Tuong Nguyen. The experimental work was supported by funds from the Bundesministerium für Bildung, Wissenschaft, Forschung und Technologie (grants BEO 0310549A, BEO 0310664, and BEO 0311203). C. Tebbe is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Young, JPW. Phylogeny and taxonomy of rhizobia. Plant and Soil 1995; 186:45-52. 2. Maier RJ, Triplett EW. Toward more productive, efficient, and competitive nitrogen-fixing symbiotic bacteria. Crit Rev Plant Sci 1996; 15:191-234. 3. Olsen PE, Rice WA, Bordeleau LM et al. Levels and identities of nonrhizobial microorganisms found in commercial legume inoculant made with nonsterile peat carrier. Can J Microbiol 1996; 42:72-75. 4. Paau AS. Improvement of Rhizobium inoculants. Appl Environ Microbiol 1989; 55:862-865. 5. Van Elsas JD, Heijnen CE. Methods for the introduction of bacteria into soil: A review. Biol Fertil Soils 1990; 10:127-133. 6. Stacey G. The Rhizobium experience. In: Halvorson O, Pramer D, Rogul M, eds. Engineered organisms in the environment: Scientific issues. Washington: ASM Press, 1985:109-120. 7. Lowendorf HS. Factors affecting the survival of Rhizobium in soil. In: Alexander M, ed. Advances in Microbial Ecology. New York: Plenum Press, Vol.4, 1977:87-124. 8. Handley BA, Hedges AJ, Beringer JE. Importance of the host plants for detecting the population diversity of Rhizobium leguminosarum biovar viciae in soil. Soil Biol Biochem 1998; 30:241-249. 9. Laguerre G, Bardin M, Amarger N. Isolation from soil of symbiotic and nonsymbiotic R. leguminosarum by DNA hybridisation. Can J Microbiol 1993; 39:1142-1149.


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10. Louvrier P, Laguerre G, Amager N. Semiselective medium for isolation of Rhizobium leguminosarum from soils. Soil Biol Biochem 1995; 27:919-924. 11. Jansson JK. Tracking genetically engineered microorganisms in nature. Current Opinion in Biotechnology 1995; 6:275-283. 12. Wilson KJ. Molecular techniques for the study of rhizobial ecology in the field. Soil Biol Biochem 1995; 27:501-514. 13. Doyle JD, Stotzky G, McClung G et al. Effects of genetically engineered microorganisms on processes in natural habitats. Advances in Applied Microbiology 1995; 40:237-287. 14. Sessitsch A, Jjemba PJ, Hardarson G et al. Measurement of the competitiveness index of Rhizobium tropici strain CIAT899 derivatives marked with the gusA gene. Soil Biol Biochem 1997; 29:1099-1110. 15. Vahjen W, Munch JC, Tebbe CC. Fate of three genetically engineered, biotechnologically important microorganisms species in soil: Impact of soil properties and intraspecies competition with nonengineered strains. Can J Microbiol 1997; 43:827-834. 16. Selbitschka W, Pühler A, Simon R. The construction of recA-deficient Rhizobium meliloti and R. leguminosarum strains marked with gusA or luc cassettes for use in risk-assessment studies. Molecular Ecology 1992; 1:9-19. 17. Selbitschka W, Arnold W, Priefer UB et al. Characterization of recA genes and recA mutants of Rhizobium meliloti and Rhizobium leguminosarum biovar viciae. Mol Gen Genet 1991; 229:86-95. 18. Hagen M, Pühler A, Selbitschka W. The persistence of bioluminescent Rhizobium meliloti strains L1 (RecA-) and L33 (RecA+) in nonsterile microcosms depends on the soil type, on the co-cultivation of the host legume alfalfa and on the presence of an indigenous R. meliloti population. Plant and Soil 1997; 188:257-266. 19. Dammann-Kalinowski T, Niemann S, Keller M et al. Characterization of two bioluminescent Rhizobium meliloti strains constructed for field releases. Appl Microbiol Biotechnol 1996; 45:509-512. 20. Garland JL, Mills AL. Classification and characterization of heterotrophic microbial communities on the basis of patterns of community-level soil-carbon-source utilization. Appl Environ Microbiol 1989; 57:2351-2359. 21. Vahjen W, Munch JC, Tebbe CC. Carbon source utilization of soil extracted microorganisms as a tool to detect the effects of soil supplemented with genetically engineered and nonengineered Corynebacterium glutamicum and a recombinant peptide at the community level. FEMS Microbiol Ecol 1995; 18:317-328. 22. Thimm T, Tebbe CC. - unpublished results 23. Schwieger F, Willke B, Munch JC et al. Ecological pre-release risk assessment of two genetically engineered, bioluminescent Rhizobium meliloti strains in soil column model systems. Biol Fertil Soils 1997; 25:340-348. 24. Schwieger F, Dammann-Kalinowski T, Dresing U et al. Environmental release of luciferase marker gene-tagged Sinorhizobium meliloti strains into lysimeters to evaluate the ecological significance of soil inoculation and a recA-mutation. FEMS Microbial Ecol–manuscript submitted 25. Dresing U, Schwieger F, Dammann-Kalinowski T et al. Field release of two bioluminescent Sinorhizobium meliloti strains to monitor survival and dissemination of a genetically engineered soil bacterium in the environment. – manuscript submitted. 26. Schwieger F, Tebbe CC. Impact of a field release of S. meliloti L33 on the structure of bacterial communities in the rhizosphere of a host and a nonhost plant as detected by nucleic acid based techniques. – manuscript submitted.

CHAPTER 10

The Field Release and Monitoring of Rhizobial Strains Marked with lacZ and Mercury Resistance Genes Viviana Corich, Alessio Giacomini, Elena Vendramin, Patrizia Vian, Milena Carlot, Andrea Squartini and Marco P. Nuti

10.1. Introduction

T

his brief account will summarize the experiences gathered over an eight year period (1989-1996), devoted to assessing the potential risks associated with the construction and use of genetically modified microorganisms. As part of the BAP, BRIDGE and IMPACT I European projects, these studies fall primarily in the category of prenormative research; the aims of which were to provide information to competent authorities and protection agencies and to contribute to the build up of case histories for the validation of environmentally safe guidelines. With this respect, the experimental schemes adopted have been focused on addressing a simple and somewhat rhetorical question: whether or not a neutral genetic modification, not conferring advantages to the bacteria, could perturb the environment into which they are released?

10.2. Designing the Markers The theoretical rationale that inspired our marker design stemmed from considerations on the strength of gene promoters. In the late 1980s we had scanned aligned compilations of bacterial promoter sequences and summarized that a hypothetical consensus promoter, featuring the most frequent base entry displayed at each position by known upstream gene regions, was likely to confer strong expression levels. We therefore constructed a 59-mer synthetic promoter, which was then assembled in a gene cartridge (Fig.10.1), together with a lacO-lacIq system to allow regulated expression, translational requirements by means of a ribosome binding site, and the lacZ reporter gene.1 In addition a number of suitable restriction sites were added allowing: (1) the subcloning of the module, (2) optional removal of the operator to attain constitutive expression and (3) insertional gene fusions replacing or trailing the lacZ gene. In order to add selectable traits for future monitoring of the GMM, mercury resistance determinants were included in the cassette. The avoidance of antibiotic markers was appropriate in the light of EU guidelines which discourage their use in open field releases. A broad host-range vector of the incQ type was the carrier of the cassette which was tested in Rhizobium leguminosarum and E. coli. The incorporation of inorganic Hg as a selective agent safeguarded against extensive fungal growth on isolation plates. The Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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Fig.10.1. Map of the assembled gene cartridge used for the genetic modifications (redrawn with modifications from ref. 1).

synthetic promoter fulfilled expectations, conferring levels of β-gal activity that more than doubled those of the tac promoter cloned in an isogenic vector-background. Interestingly, the regulated version of the cassette behaved differently in the two tested species: in E. coli an IPTG-induced regulated tac promoter was stronger than the regulated synthetic promoter while in Rhizobium the situation was reversed. The cassette proved therefore efficient for monitoring of rhizobia. An additional advantage was the facility of performing highlyspecific fragment amplification from environmental samples ensured by a DNA sequence virtually unique in nature.

10.3. Construction of GMM Strains 1110, 1111 and 1112 Our first target organism was Rhizobium leguminosarum bv. viciae strain 1003. To begin with, a spontaneous rifampicin-resistant derivative was obtained, in which the above described gene cassette was introduced (such spontaneous antibiotic resistances do not fall in the “recombinant” category regulated by the EU). In order to observe the effects that identical genetic modifications could exert on a microorganism in relation to the replicon of residence and to the status of marker gene expression, we devised and constructed three isogenic GMM derivatives of strain 1003.2 The first (strain 1110) carried the lac-regulated mercury-resistant gene cassette on a broad host range incQ plasmid derived from pRL4973 and devoid of antibiotic resistances (plasmid pDG3). The second (strain 1111) was identical with the exception of the absence of the 26 bp lac operator (plasmid pDG4). This difference brought about a constitutively high level of lacZ expression. The third strain (1112) had the same regulated cassette of strain 1110, but inserted into the chromosome via homologous recombination of flanking Rhizobium sequences previously cloned in a suicide vector. The three constructs were made to test whether the same marker genes could give rise to different outcomes in terms of stability and fitness, with respect to their positions and expression levels within the cells. Phenotypically the GMMs were clearly distinct from their parent strain due to the levels of β-gal activity and to their resistance to HgCl2 (Table 10.1). Mercury resistance was also found to be affected by the growth medium, being lower in mannitol-based media, supposedly due to an interference between mannitol metabolism and mercury detoxification.4 Measurements of growth kinetics indicated that the genetic modification did not affect growth rate of the three strains. When testing genetic stability through 70 generations in batch culture, as predicted, the strain carrying the unregulated cassette underwent an over 99% loss of the introduced plasmid, versus a 99% marker stability shown by the regulated plasmid version over the same period. The metabolic load of a continuous lacZ expression at very high levels therefore appears a deleterious burden that prompts marker/reporter elimination.

10.4. Microcosm Studies of GMMs A long-term microcosm study to reproduce storage of commercial agricultural inoculants was undertaken to obtain information on the behavior of GMMs during a mandatory stage for any marketed PGPR (Plant Growth Promoting Rhizobacteria). The collaboration with Heligenetics S.p.A (Gaiba [Rovigo] Italy), a company manufacturing inoculants, allowed us to use state-of-the-art industrial technology and test

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Table 10.1. Expression level of the introduced genes: β-galactosidase activity and resistance to mercury chloride Strain

1003 (wt) 1110 1111 1112

β-gal activity (Miller U)

β-gal activity (Miller U)

IPTG-induced

uninduced

MIC (µg/ml) of HgCl2

38 6.849 12.982 718

40 260 15.165 243

250 nodule occupants from the first release site. The Rhizobium leguminosarum bv. viciae natural local population showed more than 30 different RAPD profiles some of which were present in a large number of nodules. Nodulation tests on an alternative host, Vicia faba subsp. minor, also contributed data to identify the profile of the dominant strain which occupied over half of the nodules on both plants (profile D on Table 10.2). A spontaneous rifampicin resistant derivative of this strain was genetically modified by introducing plasmid pDG3, yielding the equivalent of strain 1110 used in the first release, but in a background strain native of the chosen habitat and successful on its symbiotic hosts. The strain was named 1114. The performance of this GMM lived up to the standards of its parent whose competitive traits were maintained in terms of nodulation and persistence in the habitat. Despite the active colonization and infection of strain 1114, again the resident microbiota did not show significant alterations, as determined by plate counting, indicating that a GMM extensively invading its environment, does not necessarily alter the measured soil populations.6 An analysis of heterospecific but related species such as Sinorhizobium meliloti was carried out from pea rhizosphere using MPN counts by means of the homologous host alfalfa. The GMM did not alter S. meliloti numbers. Moreover, their growth appeared not to be stimulated in the rhizosphere of the heterologous pea plant. In terms of marker/reporter genes efficiency in monitoring, differences were noticed between the first released GMM, based on strain 1003, and 1114, the dominant “profile D” strain. The former, despite carrying mercury resistant genes, was difficult to grow on mercury con-

Table 10.2. Frequency of the most abundant RAPD profiles shown by nodule occupants on pea (P. sativum) and vetch (V. faba) in the soil of the release P. sativum (250 nodules)

V. faba (32 nodules)

A

10%

13%

B

9%

19%

C

11%

6%

D

55%

44%

E

< 1%

9%

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taining media when plated straight from soil suspension, although it would grow well if restreaked from agar plates or liquid cultures. This is presumably due to the necessity for induction of mercury resistance. The latter strain, 1114 did not present such a limitation, allowing a faster screening. Conversely, the lacZ marker driven to high expression by the synthetic promoter, conferred a distinct phenotype in the 1003 background, while in strain 1114 a higher endogenous β-galactosidase activity made the reporter less distinctive when monitoring the GMMs.

10.7. Conclusion In essence, the experiments described here indicate that the efficiency of detoxifying and catabolic marker genes can not be generalized and is dependent on the physiology of each bacterial strain. Their stability instead, is correlated with both gene expression and genomic location. As for field release practice, data suggest that the use of an allochtonous strain does not warrant successful colonization, and that on the contrary, strong autochtonous isolates can yield backgrounds for a more promising field exploitation. GMM impact on resident life is not necessarily associated with niche occupation and will therefore be dependent on the nature of the introduced genetic modification.

Acknowledgment M.P Nuti is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Giacomini A, Ollero FJ, Squartini A et al. Construction of multipurpose gene cartridges based on a novel synthetic promoter for high-level gene expression in gram-negative bacteria. Gene 1994; 144:17-24. 2. Corich V, Bosco F, Giacomini A et al. Fate of genetically modified Rhizobium leguminosarum biovar viciae during long-term storage of commercial inoculants. J Appl Bacteriol 1996; 81:319-328. 3. Elhai J and Wolk CP. A versatile class of positive-selection vectors based on the nonviability of palindrome-containing plasmids that allow cloning into long polylinkers. Gene 1988; 68:119-138. 4. Robinson JB, and Tuovinen OH. Mechanisms of microbial resistance and detoxification of mercury and organomercury compounds: Physiological, biochemical and genetic analyses. Microbiol Rev 1984; 48:95-124. 5. Corich V, Giacomini A, Concheri G et al. Environmental impact of genetically modified Azospirillum brasilense, Pseudomonas fluorescens and Rhizobium leguminosarum released as soil/seed inoculants. In: Jones D, ed. Biosafety Results of Field Tests with Genetically Modified Plants and Microorganisms. Oakland, University of California Division of Agriculture and Natural Resources 1995: 371-388. 6. Nuti MP, Basaglia M, Bonfante P et al. Field release of genetically modified biofertilizers and phytostimulators. In: Matsui S, Miyazaki S, and Kasamo K, eds. The Biosafety Results of Field Tests of Genetically Modified Plants and Microorganisms. Tsukuba, Ibaraki, Japan: JIRCAS Publisher, 1996: 101-111.

CHAPTER 11

The Field Release and Monitoring of GUS-Marked Rhizobial Strain CT0370 Penny R. Hirsch, Tom A. Mendum, Alfred Pühler and Werner Selbitschka

11.1. Introduction

T

he population genetics of soil bacteria provides strong evidence that genes are exchanged in the natural environment (For examples, see refs 1-3). However, there is little information on the frequency and time scale of such events. There are concerns that the deliberate release of beneficial genetically modified microorganisms (GMM) as plant inoculants could result in the generation of undesirable hybrids if novel genes are transferred from the GMMs to the native population. Conversely, genes from native strains could be transferred into the inoculant, affecting its performance or persistence. There is extensive information on the survival of rhizobial inoculants,4 which have been widely used for over 100 years in agriculture to form dinitrogen-fixing symbioses with leguminous plants. In recent years there have been several releases of GM rhizobia (see refs 5-7 and Chapters 9 and 10). Also evidence has been obtained for the transfer of a conjugative transposon from an inoculant to native rhizobia,8,9 but there are no reports of genes being transferred into rhizobial inoculants. To investigate this possibility, Selbitschka et al10 marked a strain of Rhizobium leguminosarum biovar viciae which had been cured of its symbiotic plasmid and consequently had lost the ability to form nodules on pea plants, by insertion of the Escherichia coli uidA (gusA) gene into the chromosome. This gene conferred β-glucosidase (GUS) activity, not normally present in rhiziobia or plants (see Chapter 6). When the strain, CT0370, re-acquired a conjugative symbiotic plasmid, it regained nodulation ability and formed nodules with GUS activity. Thus, the GUS marker not only facilitated the monitoring of the survival of CT0370 after field release, but it also provided a simple screen: any pea root nodules containing rhizobia with GUS activity would indicate symbiotic gene acquisition by CT0370. To increase the potential for the detection of transfer events, the release site selected contained an established population, released seven years previously by Hirsch and Spokes,6 of R. leguminosarum biovar viciae strain RSM2004 containing a conjugative symbiotic plasmid that had been shown to confer nodulation ability on strain CT0370.

11.2. Construction of GMM Strain CT0370 The genetically modified release strain CT0370 was constructed as follows. The parent strain LRS39401 is a derivative of R. leguminosarum biovar viciae faba bean nodule isolate VF39 cured of its symbiotic plasmid, and selected for spontaneous mutation to streptomycin resistance.11 It was subjected to targeted insertion of the E. coli gusA gene constitutively expressed by the promoter PnptII of the nptII gene of transposon Tn5 into a noncoding Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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Tracking Genetically Engineered Microorganisms

sequence between the chromosomal recA and alaS genes, reported by Selbitschka et al12 In order to accomplish integration of the gus gene cassette, Selbitschka et al10 introduced an XhoI restriction site by site-directed mutagenesis in the recA-alaS region, immediately adjacent to the putative rho-independent terminator of the recA gene. The chromosomal site had been chosen since previous characterization of Sinorhizobium meliloti strain L33 which carried the bioluminescence mediating firefly luc gene in the analogous recA-alaS intergenic region had shown that an insertion in the target site per se did not adversely affect the strain’s fitness. Selbitschka et al13 reported that strain L33 behaved like its parent strain with respect to vegetative and symbiotic properties such as growth rate, growth competitive abilities or symbiotic performance (see also Chapter 9). The recA-alaS region was cloned in E. coli using a vector which could not replicate autonomously in Rhizobium, and contained sacRB (sucrose sensitivity) and aacC1 (gentamicin resistance) markers. Following mobilization into strain LRS39401, gentamicin resistant transconjugants were assumed to have undergone a chromosomal insertion of the vector in the target site by homologous recombination. Following selection on sucrose, some transconjugants had lost the gentamicin and sucrose-sensitivity markers of the vector but retained a GUS phenotype. These were tested for vector sequences by extracting DNA, digesting with BamH1, EcoR1 and NruI, and probing gel blots with vector DNA. PCR with primers designed to identify a single GUS gene cassette insertion in the predicted position was used to confirm the structure (Fig. 11.1). Primers were: recA forward, uidA reverse, uidA forward, and alaS reverse. PCR products arose only from reactions with recA forward, uidA reverse and uidA forwards, alaS reverse. Spontaneous mutation to spectinomycin resistance was selected in one such construct, producing strain CT0370.

11.3. Detection of Strains CT0370 and RSM2004 Routine CT0370 culturing was on complete TY or minimal Y medium, detailed by Hirsch and Skinner,14 at 28˚C. To count CT0370 in soil, selective TY or Y agar were used, containing 500 µg ml-1 streptomycin (Str) and 200 µg ml-1 spectinomycin (Sp) to counterselect other bacteria, cycloheximide 100 µg ml-1 and benomyl at 7.5 µg ml-1 to inhibit fungi, and the GUS substrate X-gluc (5-bromo-4-chloro-3-indolyl-(-D)-glucoronic cyclohexamine salt, supplied by NBL Biologicals, U.K.) at 50 µg ml-1. In soil from the release site before inoculation there were 2 x 104 Str and Sp resistant bacteria and 3% of these were GUS-positive. After release the only bacteria with a typical rhizobial morphology and which gave blue colonies on the selective agar were demonstrated to be CT0370 from plasmid profiles using a modified Eckhardt procedure.14 Strain RSM2004, which contains a conjugative symbiotic plasmid marked with the neomycin resistance transposon Tn5, could be counted on TY agar containing neomycin (Neo) 100 µg ml-1, rifampicin (Rif) 50 µg ml-1 and the antifungal agents, as reported by Hirsch and Spokes.6 At each sampling time, 10 soil sub-samples (five

Fig. 11.1. GUS gene insertion in the chromosome of strain CT0370 Restriction sites: N-NruI; E – EcoRI; B – BamHI PCR primer binding sites: rf – recA forward; ur – uidA reverse; uf – uidA forward; ar – alaS reverse

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from each section of the noninoculated buffer zones) were collected and pooled, sieved through a 5 mm mesh, 10 g resuspended in 100 ml sterile distilled water, and 100 µl plated on five replicate plates. Total nodulating rhizobia were enumerated using Vicia hirsuta infection tests with most probable number (MPN) estimation as described by Hirsch and Skinner.14 Strains CT0370 and RSM2004 could be detected in DNA extracted directly from soil using PCR with primers specific for the GUS insert and Tn5, respectively, as reported by Cullen et al.15

11.4. Inoculant Preparation CT0370 inoculant was prepared using a commercial peat-based carrier system (MicroBio Ltd, Hemel Hempstead, U.K.) as described previously.6 This system releases rhizobia into the soil over a period of 1-2 weeks. Prior to release, the inoculant was enumerated in the peat and its viability was checked using fluorescent microscopy with Baclight(TM) live/ dead stain (Molecular Probes, Oregon, USA). Prior to release, direct and plate counts of the inoculant gave 1.5 x 109 and 1.7 x 109 cfu g-1 peat, respectively.

11.5. Field Release The release site (Fig. 11.2), a 3 x 3 m plot within the site of the RSM2004 release6 had been under rotation with cereal crops since 1989. To comply with the requirements of the U.K. Department of the Environment, the plot was covered by a net to exclude birds and surrounded by two 1 m buffer zones of wheat separated by a fence.

Fig. 11.2. Release plot of strain CT0370 A – wheat field, B – outer buffer zone; C – fence; D – inner buffer zone; E – net; F – peas inoculated with strain CT0370; G – dustbin.

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The release took place in July 1994: inoculant granules were placed in drills (10 cm apart), into which 840 untreated Avola peas (PGRO, Peterbourgh, U.K) coated in inoculant were planted at 10 cm intervals. A total of 662 g of peat inoculant containing 9 x 1011 live cells was applied; initial soil samples thus contained 4.9 x 105 cfu CT0370 g-1 soil. Samples were taken immediately before and after the release, then weekly for 10 weeks and then fortnightly. Peas were again planted in May 1995 and July 1995, but without further inoculation. An initial ten-fold drop in CT0370 cfu numbers was observed during the first 10 weeks, to 5 x 104 culturable cells g-1 soil (Fig. 11.3). Subsequently numbers remained around 104, similar to the numbers of native R. leguminosarum in the soil (Fig. 11.3). The inoculant appeared to survive better than RSM2004 which had rapidly dropped to 102 cfu g-1 soil although it subsequently persisted at this level. No CT0370 was detected in any buffer zone samples: the major dispersion mechanism in arable fields is via soil cultivation6 and the buffer zones were not cultivated.

11.6. Screening for pSym Acquisition by CT0370 in the Field Pea plants were harvested at pod-fill, the roots washed in water, nodules removed and surface sterilized by washing in 70% ethanol for 30 s, rinsing in H2O, leaving in 10% sodium hypochlorite for 1 min, then washing twice in H2O. They were stored at –70˚C in sterile 96 well microtiter dishes containing sterile 10% glycerol. The nodules were screened for GUS activity using MUG (4-methyl-umbelliferyl-β-D-glucoronide supplied by NBL biologicals) and UV-induced fluorescence was assessed as described.10 Nodules containing GUS activity were crushed onto TY plates. Over two years, 21,146 nodules were screened. Three had GUS activity but were found to contain nonrhizobial contaminants. Hence, no rhizobia with GUS activity were found.

11.7. Plasmid Transfer from RSM2004 to CT0370 For filter crosses, parents were grown in liquid TY to mid-late log-phase. Then, 100 µl of each culture were placed on a membrane filter and incubated overnight at 28˚C on TY

Fig. 11.3. Survival of strain CT0370 and other rhizobia in the field


149

plates. The mixed growth was suspended in 1 ml H2O by vortexing and serial dilutions were plated onto selective media. Sterile field soil was prepared by drying and milling, passing it through a 5 mm sieve and autoclaving it three times at 121˚C for 1 hr with 24 hr intervals between sterilizations. Nonsterile conjugations were performed in fresh field soil (7.5% w/w H2O) previously passed through a 5mm sieve. Samples from late log phase cultures of CT0370 and RSM2004 were mixed, centrifuged, and resuspended in 500 µl/H2O. Then, 5 g of soil were added, the mixture was vortexed and incubated for 4 days at 28˚C. Samples of parental cultures, and of soil after 2 and 4 days (resuspended as described for field soil) were diluted and plated onto selective media. To determine how plasmid transfer rate was affected by the presence of pea plants, and of introducing inoculant in peat granules, field soil in pots was mixed with peat inoculant to give RSM2004 and CT0370 levels of approximately 106 cfu g-1 soil, and peas were planted. Control pots contained the parents alone or strain CT0370 carrying pSym2004. Samples of the inoculant granules and of rhizosphere soil after one and two weeks were resuspended and plated as described for field samples. Root nodules were screened for GUS activity as described previously. The highest transfer frequency of the pSym plasmid from RSM2004 to CT0370, i.e., 9 x 10-5 transconjugants per recipient, was observed in laboratory matings on filters. In sterile soil microcosms with 106 to 107 cfu of each parent strain per g soil, the highest frequency was 2.6 x 10-6 transconjugants per recipient cell; in nonsterile soil it was 2.3 x 10-7. In pot experiments with peat inoculant containing both parents, providing 5 x 106 cfu g-1 soil, no transconjugants were found, indicating that in field soil with only 102-103 RSM2004 and 104-105 CT0370 cfu g-1 soil, the frequency would most likely be too low to detect. CT0370 colonies reisolated from soil were screened for the acquisition of other plasmids from the soil population by subjecting them to PCR with primers designed to amplify two rhizobial plasmid replication origins identified in the field population.16,17 More than 1000 colonies were screened, pooled in groups of 10 for DNA extraction and tested by PCR. No positive results were obtained, although control reactions in which one colony of a positive field isolate was included did give the expected band.

11.8. Conclusion Although strain CT0370 lacked symbiotic genes, it survived well in field soil after release and remained at a level similar to that of the native rhizobial population for five years of sampling. This compares to the prior release of RSM2004 at the same site, where numbers declined by two orders of magnitude in the six months following release, but subsequently stabilized at around 102 cfu g soil-1.4,6 However, RSM2004 does not persist in all soils and it is probable that survival is influenced by both soil type and bacterial strain.4 The GUS marker alone would not have facilitated counting of CT0370 because of GUS activity in the indigenous population of heterotrophic soil bacteria. However, in conjunction with the chromosomal antibiotic resistances of CT0370, the gus gene made it possible to detect the strain in soil against the background population, without effecting the enumeration of RSM2004 (which contains different antibiotic resistances, neomycin and rifampicin). The GUS marker offered a very sensitive detection system for identifying root nodules with containing rhizobia with GUS activity and facilitated the screening of many more nodules than would have been feasible with markers. The lack of transconjugants with GUS activity in the field indicates that plasmid transfer does not occur at elevated levels in the field compared with laboratory studies. Symbiotic plasmid transfer from RSM2004 to CT0370 could be detected in soil microcosms with parental densities of 106-107 cfu g soil-1, much higher than those in field soil. The population density of rhizobia on pea roots is much higher than in bulk soil but did not appear to be sufficient to enable conjugation to occur at a detectable level and no evidence was found for acquisition of symbiotic or other plasmids from native

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rhizobia by CT0370. Nevertheless, studies on natural populations of rhizobia indicate that plasmid transfer does occur over time.1, 3 This contrasts with the findings by Lilley et al,2 of high levels of plasmid transfer in pseudomonad populations in the sugar beet phytosphere and indicates that rates of gene transfer in the environment vary between different bacteria groups and habitats. In conclusion, the GUS marker facilitated sensitive detection of CT0370 in root nodules and proved to be an excellent marker for monitoring microbes in the field.

Acknowledgments This work was supported in part by EC Biotechnology Programmes (BIO2 CT92-0370 and BIO4 CT96-0434). IACR-Rothamsted receives grant-aided support from the Biotechnology and Biological Sciences Research Council of the UK. A. Pühler and W. Selbitschka are members of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

References 1. Young JPW, Wexler M. Sym plasmid and chromosomal genotypes are correlated in the field populations of Rhizobium leguminosarum. J Gen Microbiol 1988; 134:2731-2739. 2. Lilley AK, Bailey MJ. The acquisition of indigenous plasmids by a genetically marked pseudomonad population colonising the phytosphere of sugar beet is related to local environmental conditions. Appl Environ Microbiol 1997; 63:1577-1583. 3. Villadas PJ, Burgos P, Rodriguez-Navarro DN et al. Characterization of rhizobia homologues of Sinorhizobium meliloti insertion elements ISRm3 and ISRm4. FEMS Microbiol Ecol 1998; 25:341-348. 4. Hirsch PR. Population dynamics of indigenous and genetically modified rhizobia in the field. New Phytol 1996; 133:159-171. 5. Bosworth AH, Williams MK, Albrecht K et al. Alfalfa yield response to inoculation with recombinant strains of Rhizobium meliloti with an extra copy of dctABD and/or modified nifA expression. Appl Environ Microbiol 1994; 60:3815-3832. 6. Hirsch PR, Spokes JD. Survival and dispersion of genetically modified rhizobia in the field and genetic interactions with native strains. FEMS Microbiol Ecol 1994; 15:147-160. 7. O’Flaherty S, Moënne-Loccoz Y, Boesten B et al. Greenhouse and field evaluations of an autoselective system based on an essential thymidylate synthase gene for improved maintenance of plasmid vectors in modified Rhizobium meliloti. Appl Environ Microbiol 1995; 61:4051-4056. 8. Sullivan JT, Patrick HN, Lowther WL et al. Nodulating strains of Rhizobium loti arise through chromosomal symbiotic gene transfer in the environment. Proc Natl Acad Sci USA 1995; 92:8985-8989. 9. Sullivan JT, Ronson CW. Evolution of rhizobia by acquisition of a 500-kb symbiosis island that integrates into a phe-tRNA gene. Proc Natl Acad Sci USA 1998; 95:5145-5149. 10. Selbitschka W, Jording D, Nieman S et al. Construction and characterisation of a Rhizobium leguminosarum biovar viciae strain designed to assess horizontal gene transfer in the environment. FEMS Microbiol Lett 1995; 128:255-263. 11. Hynes MF, McGregor NF. Two plasmids other than the nodulation plasmid are necessary for formation of nitrogen-fixing nodules by Rhizobium leguminosarum. Mol Microbiol 1990; 4:567-574. 12. Selbitschka W, Arnold W, Priefer UB et al. Characterization of recA genes and recA mutants of Rhizobium meliloti and Rhizobium leguminosarum biovar viciae. Mol Gen Genet 1991; 229:86-95. 13. Selbitschka W, Dresing U, Hagen M et al. A biological containment system for Rhizobium meliloti based on the use of recombination-deficient (recA-) strains. FEMS Microbiol Ecol 1995; 16:223-232.


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14. Hirsch PR, Skinner, F.A. The identification and classification of Rhizobium and Bradyrhizobium. In: Board RG, Jones K, Skinner FA, eds. Identification Methods in Applied and Environmental Microbiology Blackwell Scientific Publications, Oxford. 1992:45-65. 15. Cullen DW, Nicholson PS, Mendum TA et al. Monitoring genetically-modified rhizobia in field soils using the polymerase chain reaction. J Appl Bacteriol 1998; 84:1025-1034. 16. Turner SL, Rigottier-Gois L, Power RS et al. Diversity of repC plasmid-replication sequences in Rhizobium leguminosarum. Microbiology 1996; 142:1705-1713. 17. Rigottier-Gois L, Turner SLT, Young JPW et al. Distribution of repC plasmid-replication sequences among plasmids and isolates of Rhizobium leguminosarum bv. viciae from field populations. Microbiology 1998; 144:771-780.

CHAPTER 12

Regulatory Aspects Kersti Gustafsson

12.1. Introduction

M

any countries have chosen to regulate GMOs (genetically modified organisms) for deliberate release into the environment. Reasons for regulation are the lack of knowledge on the fate and behavior of GMOs and inserted genomes deliberately released into the environment and lack of experience of accompanying risks. Another reason is the concern of people in general about releases of GMOs into the environment. Some countries are of the opinion that ethical aspects of these matters are important. Regulation and control facilitate increased general knowledge concerning the deliberate release of GMOs and hopefully contribute to a safe and acceptable development of the area. This chapter deals mainly with GMM (genetically modified organisms), however, much of the regulation is developed for the whole group of GMOs. The United States has chosen to regulate GMOs under the existing legislation, while the European Union (EU) has chosen to introduce new directives. Sweden has an act regulating GMOs and the responsibility among authorities is divided according to existing legislation. Since much is dependent on the vector of the gene modification, i.e., the organism, a divided responsibility among different experts must be relevant. The use of biotechnology in industrial applications as well as for research and development purposes has increased. This is also a reason to have a divided responsibility for the control of the use of GMOs.

12.2. Historical Aspects of the Regulation of Deliberate Releases of GMOs In 1986, the OECD (Organization for Economic Cooperation and Development) Council made recommendations for applications of recombinant DNA organisms in industry, agriculture and the environment.1 It was considered that “recombinant DNA techniques have opened up new and promising possibilities in a wide range of applications and can be expected to bring considerable benefits to mankind“. It was also considered that “a common understanding of safety issues raised by recombinant DNA techniques will provide the basis for taking initial steps toward international consensus, the protection of health and the environment, the promotion of international commerce and the reduction of national barriers to trade in the field of biotechnology, that assessment of potential risks of recombinant DNA organisms for environmental or agricultural applications is less developed than the assessment of potential risks for industrial applications“. Some of the recommendations are listed in Table 12.1. In 1992, the OECD published a follow-up document concerning safety considerations for biotechnology with good developmental principles (GDP) as a guide for the design of Tracking Genetically-Engineered Microorganisms, edited by Janet K. Jansson, Jan Dirk van Elsas, Mark J. Bailey. ©2000 EUREKAH.COM.

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Table 12.1. Recommendations to OECD Member countries from the OECD council concerning safety considerations for applications of recombinant DNA organisms in industry, agriculture and the environment (sections 1 and 3 reprinted with permission from OECD publications)1 1. OECD council recommends that OECD Member countries: a) share, as free as possible, information on principles or guidelines for national regulations, on developments in risk analysis and on practical experience in risk management with a view to facilitating harmonization of approaches to recombinant DNA techniques; b) examine their existing oversight and review mechanisms to ensure that adequate review and control of the implementation of recombinant DNA techniques and applications can be achieved while avoiding any undue burdens that may hamper technological developments in this field; c) recognize, when aiming at international harmonization, that any approach to implementing guidelines should not impede future developments in recombinant DNA techniques; d) examine at both national and international levels further developments such as testing methods, equipment design, and knowledge of microbial taxonomy to facilitate data exchange and minimize trade barriers between countries. Due account should be taken of ongoing work on standards within international organizations, e.g. WHO, CEC, ISO, FAO, MSDN1 e) make special efforts to improve public understanding of the various aspects of recombinant DNA techniques; f) watch the development of recombinant DNA techniques for applications in industry, agriculture and the environment, while recognizing that for certain industrial applications, and for environmental and agricultural applications of recombinant DNA organisms, some countries may wish to have a notification scheme; g) ensure that the assessment and review procedures protect intellectual property and confidentiality interests in applications of recombinant DNA, recognizing the need for innovation while still ensuring that all necessary information is made available to assess safety. 2. OECD council recommends, with specific reference to agricultural and environmental applications, that OECD Member countries: a) use the existing considerable data on the environmental and human health effects of living organisms to guide risk assessments; b) ensure that recombinant DNA organisms are evaluated for potential risk, prior to applications in agriculture and the environment by means of an independent review of potential risks on a case-by-case basis;2 c) conduct the development of recombinant DNA organisms for agricultural or environmental applications in a stepwise fashion, moving, where appropriate, from the laboratory to the growth chamber and greenhouse, to limited field testing and finally, to large-scale field testing; d) encourage further research to improve the prediction, evaluation, and monitoring of the outcome of applications of recombinant DNA organisms. 1 World Health Organization (WHO); Commission of the European Communities (CEC);

International Standards Organization (ISO); Food and Agriculture Organization (FAO); Microbial Strains Data Network (MSDN). 2 Case-by-case means an individual review of a proposal against assessment criteria which are relevant to the particular proposal; this is not intended to imply that every case will require review by a national or other authority since various classes of proposals may be excluded.

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small-scale field research with GMOs.2 The key safety factors in determining the safety of any experiment were: i) the characteristics of the organism(s) used, including the introduced genome; ii) the characteristics of the research site and the surrounding environment; and iii) the use of appropriate experimental conditions. The characteristics of importance for the microorganism included dispersal, survival, multiplication and the potential for gene transfer. The document also considered the mode of action of the GMM, including persistence and degradation of any newly acquired toxic metabolite. Additional important characteristics of the GMM, including interactions with other species and/or biological systems, were also considered. In 1995, the OECD initiated an Expert Group to manage the implementation of OECD’s Programme for the Harmonisation of Regulatory Oversight in Biotechnology. In 1998, the group was renamed as a Working Group. The Group is currently made up of experts involved in the regulation of biotechnology who have been nominated by their Member State. The Working Group is a subsidiary body to the Joint Meeting of the Chemicals Group and Management Committee of the Special Programme on the Control of Chemicals within the OECD. The main focus of the Work Programme is the international harmonization of regulatory oversight in biotechnology which will ensure that environmental health and safety aspects are properly evaluated, while avoiding nontariff trade barriers to products of the technology. The main areas of work are: i) the development of Consensus Documents on specific scientific issues related to biotechnology; ii) outreach activities, including the development and maintenance of “Biotrack Online“, that makes information on the Harmonisation Programme available to anyone interested; and iii) general issues associated with harmonization of biotechnology regulation.a The regulatory developments in Member States can be found via Biotrack Online.b

12.3. The European Union and the European Free Trade Association The European Union has chosen to regulate GMOs mainly via a directive on the contained use of genetically modified microorganisms3 and a directive on the deliberate release into the environment of genetically modified organisms.4 The directives were modeled on the chemicals notification directives.c These GMO directives also have impact on the regulation of GMMs within the European Free Trade Association (EFTA). The responsible institution for Horizontal Legislation concerning GMOs within the EU is Directorate-General XI: Environment, Nuclear Safety and Civil Protection.d For specific product legislation, the responsibility is shared by other Directorates as follows: DG III (Industry) and DG VI (Agriculture). Other Directorates involved in regulation of GMOs are DG VII (Transport)—responsible for the safe transport of GMO, and DG XII (Science, Research and Development) and The European Commission Joint Research Centre (Institute for Systems, Informatics and Safety) are responsible for information on research and development of GMOs. Apart from the above mentioned directives, other relevant regulations concern the protection of workers from the risks of exposure to biological agents (Council Directives 90/679/EEC and 93/88/EEC). For some products, there are other relevant legislations: for additives in feeding stuffs (Council Directive 93/114/EEC); for medicinal products (Council

a

http://www.oecd.org/ehs/service.htm http://www.oecd.org/ehs/country.htm c http://europa.eu.int/en/comm/dg11/guide/part2g.htm d http://www.oecd.org/ehs/cecreg.htm b

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Directive 93/41/EEC); and for novel food (European Parliament and Council’s Regulation No 258/97). Directive 90/219/EEC lays down common measures for the contained use of GMMs with a view to protecting human health and the environment. Directive 90/219/EEC was amended in October 1998.5 The objective of Directive 90/220/EEC is to approximate the laws, regulations and administrative provisions of the EU Member States and to protect human health and the environment: • when carrying out deliberate releases of GMOs into the environment for research and development purposes - Part B; • when placing products on the market that contain, or consist of, GMOs intended for subsequent deliberate release into the environment—Part C. Since disparity between rules concerning GMOs in the Member States may cause unequal conditions of competition or trade barriers it is necessary to approximate the laws in the different Member States. Although the objective of the directive is the establishment and maintenance of the internal market, nevertheless health and environmental safety aspects, as well as consumer protection should be at a high level of consideration. Before a deliberate release of a GMO into the environment is performed, the responsible person must submit a notification to the competent authority in the EU Member State where the release is going to take place. Such research and development experiments are notified according to Part B of directive 90/220/EEC. The notification must also be circulated among other Member States and EFTA states via the European Commission (further on referred to as the Commission). All countries are allowed to comment on the circulated summary notification information format (SNIF), although the comments from other states do not have a decisive effect on the decision. It has to be noted that the competent national authority takes its decision independently. The authority giving the consent must have evaluated the information and have made itself sure that the release will be safe for human health and the environment. A company that wants to place a product containing a GMM(s) on the market has to notify the product. The notification can be sent to a competent authority in any of the EU Member States before the product is for the first time placed on the EU market. Products are notified under Part C of directive 90/220/EEC. The notification must be circulated among other Member States and EFTA states via the Commission. All countries can comment on the circulated summary notification information format (SNIF). A joint decision is thereafter taken among EU Member States according to specified rules in the directive and the decision taken will be valid in all EU Member States and EFTA states. The Commission has now presented a proposal to the European Council concerning amendments of directive 90/220/EEC.6 Some main improvements suggested include: i. simplified administrative procedures; ii. the possibility for the Commission to consult a scientific committee concerning ethical consequences of biotechnology; iii. mutual principles on risk assessment; iv. monitoring of products placed on the market; and v. marketing approvals for a fixed time period. The Economic and Social Committee of the European Commission has given its diversified opinion on the proposal.7

12.4. Canada The Canadian Environmental Protection Act (CEPA) requires that all new substances, including those that are living organisms, are to be assessed for their potential to harm the e

http://www.ec.gc.ca/cceb1/eng/97brochuree.html

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environment, the environment on which life depends, or human life or health, prior to being imported or manufactured in Canada.e The CEPA New Substances Notifications (NSN) Regulations describe the information that must be provided to Environment Canada in order for this assessment to be done. The NSN regulations covering substances like chemicals or polymers, have been in effect since 1994 and the 1997 amendment covers substances that are organisms, or products of microorganisms (biochemicals and biopolymers). The amendment was published in the Canada Gazette, Part II, on March 5, 1997 and came into effect on September 1, 1997.

12.5. Switzerland On July 1, 1997, a series of amendments to the Federal Law on Environmental Protection and to the Federal Law on Epidemics were enforced concerning the use of organisms used in modern biotechnology.f The amendments provide the legal basis for specific regulations of environmental health and safety issues associated with GMOs. The new legislation covers all types of organisms and applications on the basis of the self-responsibility of the user. For the environmental use of GMOs and/or pathogenic organisms, special provisions, including risk assessment and notification or authorization procedures are requested. The legislation can be applied directly. However, regulations are needed to fully establish the new procedures and to define the role and responsibilities of the various authorities. A regulation has been proposed by the government on the use of organisms in the environment. The regulation is harmonized with the corresponding directive of the European Union 90/220/EEC.

12.6. United States The responsible agency for regulatory developments in biotechnology in the US is the United States Environment Protection Agency (EPA) concerning microbial/plant pesticides, new uses of existing pesticides and novel microorganisms.g The relevant laws are the Federal Insecticide, Fungicide, and Rodenticide Act (FIFRA) and the Toxic Substances Control Act (TSCA). In 1986, the US Office of Science and Technology Policy published a co-ordinated framework for regulation of biotechnology in the Federal Register of June 26. Within the policy it was noted which regulations could be amended to include GMMs. Microbial pesticides including GMMs are currently registered under FIFRA. On September 1, 1994, the final rule on Microbial Pesticides; Experimental Use Permits (EUP) and Notifications under FIFRA was published in the Federal Register. These regulations clarify the circumstances under which an EUP is presumed not to be required and implement a screening procedure that requires notification to the EPA before initiation of small-scale field testing of certain microbial pesticides. The EPA believes that small-scale tests in the environment with some microbial pesticides may pose sufficiently different risk considerations from conventional chemical pesticides that a closer evaluation at the small-scale testing stage may be warranted. Any person who plans to conduct small-scale testing of a microbial pesticide must submit a notification to the EPA and obtain approval for small-scale field testing involving microbial pesticides whose pesticidal properties have been imparted or enhanced by the introduction of genetic material that has been deliberately modified. The regulation under which the TSCA Biotechnology Program functions is titled “Microbial Products of Biotechnology; Final Regulation Under the Toxic Substances Control f

http://www.oecd.org/ehs/swireg.htm http://www.oecd.org/ehs/usareg.htm h http://www.epa.gov/opptintr/biotech/biorule.htm g

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Act; Final Rule” and was published in the Federal Register on April 11, 1997. An Internet website has been created to allow more efficient public, governmental and educational access to the TSCA Biotechnology Program.h The final regulation of microbial products of biotechnology under the TSCA regard commercial biotechnology research and development activities using intergeneric microorganisms for e.g., biofertilizers, biosensors, biotechnology reagents, commodity or specialty chemical production, energy applications, waste treatment or pollutant degradation and commercial biotechnology products. Persons that intend to use intergeneric microorganisms (microorganisms formed by combining genetic material from organisms in different genera) for commercial purposes in the United States should submit a Microbial Commercial Activity Notice (MCAN) to the EPA at least 90 days before such use. To test new microorganisms for commercial research and development purposes in the environment, a TSCA Experimental Release Application (TERA) should be submitted to the EPA 60 days prior to initiating field trials. New microorganisms include intergeneric microorganisms. The EPA believes that intergeneric microorganisms have a sufficiently high likelihood of expressing new traits or new combinations of traits to be termed “new“ and warrant review. Microorganisms that are not intergeneric would therefore not be “new“ and thus would not be subject to reporting under Section 5 of TSCA.

12.7. Risk Assessment

In 1993, a comparison was made between some selected risk assessment strategies.8 The necessary information generally demanded for an ecological risk assessment could be generalized as: ● Biology of the organism and characteristics of the genome ● Fate of the organism and the genome in the ecosystem ● Impact of the organism and the genome on the ecosystem ● Effects of the organism and the genome on other organisms in the ecosystem By including humans as organisms in the ecosystems, the above generalizations are also valid for effects and impact on humans. With regard to chemicals, the discussions on risks and assessment of risk started mainly after disasters had started to occur. With regard to radiation, the discussions on risks have occurred basically parallel with the development of the technique. However, with modern biotechnology and the use of GMOs, the risk discussions were initiated before the full impact of the technique was known. Therefore the discussions on risks of GMOs sometimes lack connection to real examples and technique development within the area of research. The development might actually be slowed down due to the pro-active risk discussions. With continuous risk discussions during the notification process of field trials and GMO products, the research and development rate might be slowed down. However, obvious risk aspects can easier be discovered due to the risk discussions and should be managed early in the research and development process, and this will be of benefit for the companies, the scientists and the public. In October 1998, a workshop was held in Stockholm, Sweden, which focussed on the scientific basis for risk assessment of microbiological plant protection products charged by the European Commission, DG VI.i,9 There was a general apprehension that risk assessment of GMMs and risk assessment of microbiological plant protection products have very much in common. Microorganisms intended for deliberate release are regulated in different ways according to the purpose of the release; nevertheless, the risk assessments should benefit from general multidisciplinary discussions.

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12.8. Concern About the Insertion of Genes Coding for Antibiotic Resistance There is a general concern about the increase of antibiotic resistant pathogenic bacteria. In that perspective, the introduction of antibiotic resistance selectable marker genes in GMOs is worrying. So far, most concern has referred to agricultural crops due to the risk for horizontal gene transfer, even though that risk is likely to be small or theoretical.10 The risk, even though small, is probably more realistic with GMMs. The significance for gene transfer of antibiotic resistance genes is very difficult to determine. The risk includes the transfer, the expression of the gene and the consequences followed from that event. In general, Swedish authorities involved in GMO regulation are of the opinion that the risks with using antibiotic resistance as marker genes are very small.11 Nevertheless, due to the following arguments Swedish authorities recommend a development to the use of alternative marker genes: ● alternative marker genes are available and new marker genes are being developed (see for example chapters 5-7 in this book); a health and environmental risk assessment has to be done for those as well; ● techniques exist and are being developed to remove unwanted genes, such as antibiotic resistance genes, before placing GMOs on the market; ● in some countries, such as Sweden, there is a restrictive use of antibiotics and subsequently a relatively low occurrence of antibiotic resistant bacteria; ● there is a strongly expressed consumer worry towards antibiotic resistance marker genes. Alternative markers for GMMs, such as luxAB, luc and gfp have been reported to be promising in a Norwegian report (see also chapters 5, 7, 9 in this book).12 However, it is stated that the most optimal is probably to use a combination of different marker genes. During the above mentioned Stockholm workshop, there was agreement on some aspects, one of which concerned the production of antibiosis substances by microorganisms for use in plant protection.9 “Many microorganisms produce some antibiosis substances. Interference with the use of antibiotics in human and veterinary medicine must be avoided.” The production of antibiosis substances is not the same as the use of antibiotic resistance marker genes. However, there is a small risk that antibiotic resistance marker genes could interfere with the use of antibiotics in human and veterinary medicine, and this must be avoided.

12.9. Concern about the Insertion of Genes Coding for Mercury Resistance Mercury resistance is sometimes used for selection of GMMs. However, mercury is a very toxic compound and also one of the elements which cause waste problems. The use of mercury to detect the mer operon can be avoided by e.g., using PCR (polymerase chain reaction). The use of mercury for detecting mercury resistant bacteria is not regulated. However, many countries make efforts to diminish the use of mercury in general, e.g., Sweden has regulations (SFS 1998:944 and KIFS 1998:8) on commodities containing mercury.i

12.10. Concern on the Use of Hazardous Chemicals in Connection with the Deliberate Release of a GMM The use of hazardous chemicals in relation to GMMs raises concern. Concerns and risk assessments of GMOs tend to be quite broad, which must be due to the accumulated i

http://www.kemi.se

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knowledge on risks from other areas. GMMs must often be used in connection with chemicals, e.g., as above due to selectable marker genes or for post-release cleanup. Methyl bromide has been suggested for post-release clean up. The use of very hazardous chemicals should in general be avoided in post-release treatment after field experiments.

12.11. Ethical Aspects

The proposal for a new EU-directive on the deliberate release of GMOs6 opens up the possibility for the Commission to consult scientific committees concerning ethical consequences of biotechnology. In the scope of the Swedish legislation, it is claimed that ethical concerns shall be taken into account in assessments of deliberate releases of GMMs. There seems to be a general public concern about GMOs, but the concern is dependent on the product and its usefulness. There is, in general, little concern about improved medicinal drugs, the potential benefit is obvious (for example recombinant insulin). However, there is generally a lot of concern with GMO food and a lack of usefulness is often claimed. It is difficult to foresee how GMM products for biodegradation etc. will be received. The pros and cons for the public, for the companies and/or for science, of using GMMs for different purposes must be weighed against the use of other technologies. In 1998, the Swiss voted on an initiative concerning the prohibition of the use of transgenic animals, of the release of genetically engineered plants into the environment and of patenting of plants and animals. The result was not in favor of the prohibition, however the matter illustrates some major concerns about biotechnology.

12.12. Information on Field Releases and Products Scientists, industries and authorities seem to agree that openness on deliberate releases of GMMs is of benefit for the development of the area. Therefore, it is possible to get information on releases via OECDs database “Biotrack Online“ on field releases and also on products.j,k Information on deliberate field trials notified under part B of Directive 90/220/ EEC is available via the Internet and has been summarized in Table 12.2.l However, so far there have not been many field releases with GMMs nor products composed by GMMs. Information concerning research in this area is provided via European Commission DG XII.m Examples of European field releases are given in Chapters 8-11 in this book.

Acknowledgment K. Gustafsson is a member of the MAREP Concerted Action sponsored by the European Commission Biotechnology Programme, DGXII.

j

http://www.olis.oecd.org/biotrack.nsf http://www.olis.oecd.org/bioprod.nsf l http://biotech.jrc.it/GMO.htm m http://www.cordis.lu/biotech/src/projects.htm k

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Table 12.2. Summary of deliberate field trials in the European Union notified under part B of Directive 90/220/EEC 21 October 1991—15 March 19991 47 releases with 7 bacterial genera, 1 yeast and 2 viruses have been performed in 7 EU Member States Country

GMM

Year

Main trait/Purpose

Finland

Streptococcus sp

1995

France

Bacillus sp. Bacillus sp. Pseudomonas sp. Rhizobium sp.

1993 1993 1992, 1994 1995

bioluminescence for monitoring antibiotic residues in milk bioinsecticide gene stability testing biocontrol of tomato bacterial wilt marker system

Germany

Rhizobium sp.

1994, 1997

marker system

Italy

Azospirillum sp.

gene interaction; gene stability testing

Pseudomonas sp. Pseudomonas sp. Pseudomonas sp.

1994, 1995, 1998, 1999 1994, 1995 1997 1998

Rhizobium sp. Rhizobium sp.

1994 1994, 1995

Escherichia coli Pseudomonas sp. Pseudomonas sp.

1994 1994, 1995 1998

Pseudomonas sp. Rhizobium sp. Rhizobium sp. Rhizobium sp. Saccharomyces cerevisiae Saccharomyces cerevisiae Sinorhizobium sp.

1998 1997 1997 1998 1997

biological containment; reduced survival biological containment; reduced survival fungal resistance; fungicide synthesis; phluoro-glucynol synthesis survival, persistence and dispersal increase of nodulation competitiveness interaction studies hydrogenase synthesis alpha-amylase secretion

1997

lactose metabolism

1998

increase of nodulation competitiveness

The Pseudomonas sp. Netherlands Pseudomonas sp.

1994 1996

Pseudomonas sp.

1998

marker system chitinase synthesis; phenazine-1carboxylic acid synthesis 2,4 diacetyl-phloroglucinol synthesis; phenazine-1-carboxylic acid synthesis

Autographa californica nuclear polyhedrosis virus

1993, 1994

efficacy

Bacteriophage M13 Pseudomonas sp. Rhizobium sp.

1994 1993, 1994 1993, 1994

pollution tracing survival, persistence and dispersal marker system

Spain

United Kingdom

gene interaction; gene stability testing interaction studies fungal resistance; fungicide synthesis; phluoro-glucynol synthesis gene detection; gene stability testing interaction studies

162


References 1. Organisation for Economic Co-operation and Development. Recombinant DNA Safety Considerations. 1986; 69. Paris: OECD Publications. 2. Organisation for Economic Co-operation and Development. Safety considerations for biotechnology. 1992; 50. Paris: OECD Publications. 3. The Council of the European Communities. Council Directive of 23 April 1990 on the contained use of genetically modified microorganisms (90/219/EEC). Official Journal of the European Communities 1990; L(117):1-14. 4. The Council of the European Communities. Council Directive of 23 April 1990 on the deliberate release into the environment of genetically modified organisms (90/220/EEC). Official Journal of the European Communities 1990; L(117):15-27. 5. The Council of the European Union. Council Directive 98/81/EC of 26 October 1998 amending Directive 90/219/EEC on the contained use of genetically modified microorganisms. Official Journal of the European Communities 1998; L(330):13-31. 6. Commission of the European Communities. Proposal for a European Parliament and Council Directive amending Directive 90/220/EEC on the deliberate release into the environment of genetically. Official Journal of the European Communities 1998; C(139):1-24. 7. Economic and Social Committee. Opinion of the Economic and Social Committee on the “Proposal for a European Parliament and Council Directive amending Directive 90/220/ EEC on the deliberate release into the environment of genetically modified organisms. Official Journal of the European Communities 1998; C(407):1-6. 8. Gustafsson K, Jansson JK. Ecological risk assessment of the deliberate release of genetically modified microorganisms. Ambio 1993; 22(4):236-242. 9. KemI. Proceedings “Microbiological Plant Protection Products - Workshop on the Scientific Basis for Risk Assessment”. 1999:1-77. Swedish National Chemicals Inspectorate, Solna. 10. Nielsen KM et al. Horizontal gene transfer from transgenic plants to terrestrial bacteria— a rare event? FEMS Microbiology Reviews 1998; 22:79-103. 11. Ljungquist S, Andrén R, Fermér C. Risk att resistens mot antibiotika sprids från GMO? 1999; 99(1):1. 25. Kemikalieinspektionen, Stockholm. 12. Kruse H, Jansson J. The use of antibiotic resistance genes as marker. 1997; 97(3):1-47. Norwegian Pollution Control Authority, Oslo, Norway.

Index A

G

Antibiotic resistance 17, 18, 20, 22-24, 159 Antibiotics 17, 18, 20-24 Azospirillum 87, 93-97

Gene transfer 17, 22, 24, 150 Genetically modified microorganisms 128, 155 GFP 101, 102, 104, 105, 110-112 Gfp 13, 14 Gfp gene 104, 105, 110, 112 GFP mutant 102-109 GMMs 128, 130-136, 155-160 GMO 153, 155-160 GUS 87-91, 93, 94, 96, 97, 145-150 GusA gene 87-89, 91-94, 97

B β-galactosidase 141, 144 β-glucuronidase 87-90, 94 Biocontrol 123 Bioluminescence 70, 72, 75, 81 Biomarker 101, 102, 104-106, 112 Bioreporter 101, 102, 104, 106, 111, 112 Biosensors 80, 81, 85 Biotechnology 153, 155-158, 160

H Heavy metal sensor 81

C Cell extraction 29, 30, 32, 34, 35, 39 Confocal microscopy 102, 110, 111 Cross-resistance 21 Culturable cells 1, 5, 6, 9

I Intrinsic markers 53, 63, 64

K

D

Kanamycin 20-22, 24

Deliberate release 153, 155, 156, 158-160 DNA 30, 32, 33, 35-45 DNA extraction 30, 32, 39, 44 Dormant cells 14

L

E Environmental monitoring 29 European Commission 155, 156, 158, 160

F Field lysimeters 129-134 Field release 117, 118, 122, 123, 128, 129, 133, 135, 136, 139, 143-145, 147 Field trials 141, 143, 158, 160, 161 Fingerprinting method 54, 55, 64

LacZ 139-141, 144 Luc gene 73-78, 129 Luciferase 69-78, 80, 81, 128, 134 Luminescence 69, 71, 73-82 Luminometry 75, 76, 79 LuxAB 72, 76

M Mercury resistance 139-141, 144 Metabolically active cells 1, 13

N Neomycin 18, 21-24 Nucleic acid 29, 30, 32, 33, 35, 37-39, 42-44

164

O OECD 153-155, 160

P PCR amplification 37, 38, 40, 42-44 Plant-pathogenic bacteria 96 Plasmid transfer 148-150 Promoter probe transposon 89, 91 Pseudomonas fluorescens SBW25 117

Tracking Genetically-Engineered Microorganisms Rhizobium 56-58, 60, 61, 78 Rhizobium leguminosarum 139, 140, 143, 145 Risk assessment 154, 156-159 RNA 32, 33, 35-37, 39-42, 44, 45 Root colonization 96 Rothamsted 150 RT-PCR 62, 64

S

Quantitative PCR 41

Sinorhizobium meliloti 128, 130 Soil columns 129-135 Specific hybridization probe 54, 56, 57 Subtraction hybridization 55, 58, 60 Sugar beet 117, 118, 119, 120, 121, 122

R

T

Regulation 153-159 REP-PCR 61 Reporter gene 139, 143 Reporter system 87, 89, 90, 93 Resuscitation 5-8 Rhizobia 127, 128, 132, 135, 140, 141, 143, 145-150

Thiostrepton 18, 20-23

Q

V V. vulnificus 1 VBNC (viable but nonculturable) 1-10, 12-14

Tracking Genetically-Engineered Microorganisms

Tracking Genetically-Engineered Microorganisms (Biotechnology Intelligence Unit 2)

Uncultivated Microorganisms

Uncultivated Microorganisms

Tracking Perception

Video tracking

Try Tracking!: The Puppy Tracking Primer

Electroporation Protocols for Microorganisms

Biocommunication in Soil Microorganisms

Electroporation Protocols for Microorganisms

Electroporation Protocols for Microorganisms

Electroporation Protocols for Microorganisms

Food spoilage microorganisms

Molecular Epidemiology of Microorganisms

Microorganisms and Bioterrorism

Uncultivated Microorganisms (Microbiology Monographs)

Modelling Microorganisms in Food

Modelling Microorganisms in Food

Biomedical image analysis: Tracking

Honeypots: Tracking Hackers

Gene Expression in Recombinant Microorganisms

Halophilic Microorganisms and Their Environments

Real-time multi-object tracking

Video Tracking: Theory and Practice

Radio Interferometry and Satellite Tracking

Microorganisms and Bioterrorism (Infectious Agents and Pathogenesis)

Microorganisms in Sustainable Agriculture and Biotechnology

Iron Nutrition in Plants and Rhizospheric Microorganisms

Tracking Genetically-Engineered Microorganisms