Contributors R Amann Max Planck Institute for Marine Microbiology, Celsiusstrasse 1, D-28359 Bremen, Germany F Azam Marine Biology Research Division 0202, Scripps Institution of Oceanography, University of California, San Diego, La Jolla, CA 92093, USA P
Bissett Florida Environmental Research Institute, 4807 Bayshore Blvd., Suite 104, Tampa, FL 33611, USA
KM Bjorkman Department of Oceanography, School of Ocean and Earth Science and Technology, University of Hawaii, Honolulu, HI 96822, USA J Bowman Australian Research Fellow, School of Agricultural Science, University of Tasmania, GPO Box 252-54, Hobart, Tasmania 7001, Australia DK Button University of Alaska, Institute of Marine Science, PO Box 757220, Fairbanks, AK 997755-7220, USA L Campbell Department of Oceanography, Texas A&M University, College Station, TX 77843-3146, USA D Capone Department of Biological Sciences and Wrigley Institute Environmental Studies, University of California, AHF 108, Los Angeles, CA 90089-0371, USA DA Caron University of Southern California, Department of Biological Sciences, 3616 Trousdale Pkwy, AHF 30l, Los Angeles, CA 90089-0371, USA F Chen Center of Marine Biotechnology, University of Maryland, 701 East Pratt Street, Suite 236, Baltimore, Maryland 21202, USA MF Crowley Coastal Ocean Observation Laboratory, Institute of Marine and Coastal Sciences, Rutgers University, New Brunswick, New Jersey, USA P del Giorgio Horn Point Laboratory, University of Maryland, Center for Environmental Science, P. O. Box 775, Cambridge, MD 21613, USA JE Dore Department of Oceanography, School of Ocean and Earth Science and Technology, University of Hawaii, Honolulu, HI 96822, USA J Fell University of Miami, RSMAS, Key Biscayne, 4600 Rickenbacker Cswy, Miami, FL 33149, USA Frischer Skidaway Institute of Oceanography, 10 Ocean Science Drive, Savannah, GA 31411, USA
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FO Glockner Max Planck Institute for Marine Microbiology, Celsiusstrasse 1, D28359, Bremen, Germany D G6tz Portland State University, Department of Environmental Biology, 1719 SW 10th Avenue, Portland, Oregon 97201, USA D Griffin Department of Marine Science, University of South Florida, 140 7th Ave S, St. Petersburg, FL 33701, USA RE Hodson Department of Marine Sciences, University of Georgia, Athens, GA 306022206, USA M H Howard-Jones
Skidaway Institute of Oceanography, 10 Ocean Science Drive, Savannah, GA 31411, USA
WH Jeffrey Center for Environmental Diagnostics and Bioremediation, University of West Florida, 11000 University Parkway, Pensacola, FL 32514, USA SC Jiang Environmental Analysis and Design, University of California, Irving CA 92697-7070, USA DM Karl Department of Oceanography, School of Ocean and Earth Science and Technology, University of Hawaii, Honolulu, HI 96822, USA PF Kemp G King
MSRC,SUNY Stony Brook, Stony Brook, NY 11794, USA Darling Marine Center, University of Maine, Walpole, ME 04573, USA
D Kirchman College of Marine Studies, University of Delaware, Lewes, DE 19958, USA E Lipp University of Maryland, Centre of Marine Biotechnology, 710 E Pratt Street, Baltimore, MD 21202, USA J Lukasik Biological Consulting Services of North Florida, Inc, 3330 NW 25th Ave., Gainesville, FL 32605, USA D Mitchell University of Texas, MD Anderson Cancer Center, Department of Carcinogenesis, Science Park - Research Division, Smithville, TX 78957, USA C Mobley Sequoia Scientific, Inc., Redmond, Washington, USA M Moline Biological Sciences Department, California, Polytechnic State University, San Luis Obispo, California, USA JP Montoya School of Biology, Georgia Institute of Technology, Atlanta, GA 303320230 CL Moyer
Biology Department, BI 409, Western Washington University, Bellingham, WA 98225-9160, USA
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G Muyzer Netherlands Institute for Sea Research (NIOZ), P.O. Box 59, N1-1790 AB Den Burg (Texel), The Netherlands SY Newell
Marine Institute, University of Georgia, Sapelo Island, GA 31327, USA O
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R Noble Southern California Coastal Water Research Project, 7171 Fenwick Lane, Westminster, CA 92683, USA
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JH Paul Department of Marine Science, University of South Florida, 140 7th Ave. S., St. Petersburg, FL 33701, USA J Pernthaler Max Planck Institute for Marine Microbiology, Celsiusstrasse 1, D-28359 Bremen, Germany L Poorvin Department of Microbiology, University of Tennessee, M409 Walkers, Knoxville, TN 37996, USA P Quang University of Alaska, Department of Mathematical Sciences, PO Box 757220, Fairbanks, AK 997755-7220, USA A-L Reysenbach Portland State University, Department of Environmental Biology, 1719 SW 10th Avenue, Portland, Oregon 97201, USA BR Robertson University of Alaska, Institute of Marine Science, PO Box 757220, Fairbanks, AK 997755-7220, USA J Rose Department of Marine Science, University of South Florida, 140 7th Ave S, St. Petersburg, FL 33701, USA H Schafer Netherlands Institute for Sea Research (NIOZ), P.O. Box 59, N1-1790 AB Den Burg (Texel), The Netherlands O Schofield Coastal Ocean Observation Laboratory, Institute of Marine and Coastal Sciences, Rutgers University, New Brunswick, New Jersey, USA W Schonhuber Institute Pasteur, Physiologie Microbienne, 28 Rue du Docteur Roux 75724, Paris, Cedex 15, France B Sherr
COAS-OSU, 104 Ocean Admin. Bldg., Corvallis, OR 97331-5503, USA
E Sherr
COAS-OSU, 104 Ocean Admin. Bldg., Corvallis, OR 97331-5503, USA
G Steward Monterey Bay Aquarium Research Institute, 7700 Sandholdt Road, Moss Landing, CA 95039-0628, USA PJ Turner Ocean Sciences Department, University of California, Santa Cruz, 110 High St., Santa Cruz, CA 95064, USA PG Verity Skidaway Institute of Oceanography, 10 Ocean Science Drive, Savannah, GA 31411, USA
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SW Wilhelm Department of Microbiology, University of Tennessee, M409 Walkers, Knoxville, TN 37996, USA A Yayanos University of California, San Diego, Scripps Institution of Oceanography, 3115 Hubbs Hall, 9500 Gilman Dr.; Department 0202, La Jolla, CA 92093-0202, USA JP Zehr Ocean Sciences Department, University of California, Santa Cruz, 110 High St., Santa Cruz, CA 95064, USA
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Preface The field of marine microbiology is in the midst of a revolution. The institution of molecular methods in our field has enabled unimaginable breakthroughs in the understanding of the activities and tremendous diversity that encompasses the microbes of the oceans. My purpose in undertaking this book at this time was to bring together an up-to-date collection of works that deal with what I consider the central themes in marine microbiology as well as areas that show the breadth of the field. In addition to papers that deal with the central issues of microbial abundance, activity, and diversity, I have also included chapters on detection of phytoplankton by remote sensing (Chapter 26), marine pollution microbiology (Chapters 27 and 28), and microbes in extreme environments (Chapters 2% 30 and 31). Prior works on methods have been published under the umbrella of aquatic microbial ecology, while this book focuses on problems central to marine microbiology. After all, nearly all modern methods used today in environmental microbiology (and some even in clinical microbiology) were either developed or first applied by marine microbiologists. This includes measurement of microbial activity in situ, the detection and identification of phylotypes present by amplification, probing a n d / o r sequencing, and many other techniques. The issues which are central to marine microbiology are reflected in all the chapters, but are perhaps best conveyed by the historical perspective of Farooq Azam (The 'methods' in our madness) and the constraints put on us by sampling our huge culture, the ocean (Karl's chapter (2) on sampling). The latter chapter clearly shows that processes in the oceans occur on timescales larger than the average duration of a research grant (3 years)! 1 have also avoided topics such as bioremediation, which are of paramount importance in terrestrial and groundwater microbiology, but only have peripheral application to the marine environment. The intent of this book is to be a complete compendium of methods which will be of value to established investigators in marine microbiology, and also as a primer for those about to enter this field (graduate students and post-doctorals, primarily). Additionally, such a collection will be a useful resource to our colleagues in other disciplines in the ocean, aquatic, and environmental sciences, so that they might get a glimpse of the 'Methods' to our madness!
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1 Introduction, History, and Overview: The 'Methods' to Our Madness
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Revolutionary discoveries in marine microbiology during the past two decades have fundamentally changed our concepts of the structure and functioning of marine ecosystems. New optical, radiotracer and molecular methods showed that earlier methods had missed most of the microbial biomass, activity and diversity, and that microbes constitute the dominant biological force in shaping the ocean's biogeochemical dynamics. As a result of these discoveries, marine microbiology, long considered a fringe discipline, has emerged as arguably the most exciting and dynamic field of oceanography. No serious study of marine ecosystems today would exclude microbial processes from its scope. This is a good time, then, to look back, as I do here, to see where we have been and to examine the relationship between methods and conceptual advances in marine microbiology. We will see that the journey has been a remarkable one. Is it not remarkable that for a hundred years we could sustain the intensity of our seemingly hopeless pursuit to fathom the intricate lives of microbes in the vast seas while equipped with little more than nutrient agar and the microscope! What madness! In recent years, the methodology has become sophisticated, powerful and diverse as reflected in this book. An historical perspective should also be instructive in pondering the future. I will argue that future progress will critically depend on methods which explicitly treat the microbe's environment as an integral and indispensable part of the inquiry and hence enable us to integrate microbial processes into the structure and dynamics of marine ecosystems. It is axiomatic among its practitioners that marine microbiology is chronically 'method-limited'. But, surely, marine microbiology is not uniquely so. General, clinical and soil microbiology have also depended on methods keeping pace with emerging questions (and the methods of these fields have been freely available to marine microbiologists). Arguably, the central issues of marine microbiology are so complex that methods to explore them have been lacking. This is particularly so since the enviromnent is an integral part of the central questions, such as: what microbes inhabit the varied environments of the sea, what are their adaptive strategies for performance and persistence in their environments and how does the playing out of their adaptive strategies in the ocean ME-I'I IODS IN MICROBIOLO(;Y, VOLUME 30 ISBN0 12 521530 4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any f o r m r e s c r \ c d
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integrate into the structure and behavior of marine ecosystems? In order to address these questions, historically, the mismatch between the need and the capability of the methods has indeed been great. We could not even observe and count most microbes in marine samples until relatively recently, let alone elucidate their adaptive biology and ecosystem functions. What have been the methodological challenges in unraveling this complexity? What successes and failures to address the foregoing questions have led us to the present state of the art in methodology? What can we learn from our experience with past and current methods as we think of the methodology of the future? I will address these question in this chapter. Early microbial oceanographers used the viable count and pure culture technique to determine the abundance, distribution and species composition of bacteria. Bernhard Fischer, a professor of hygiene in Kiel, Germany, was the first to show that indigenous bacteria existed on the high seas (Fischer, 1894). He used a modified nutrient agar method (substituting beef broth in Robert Koch's recipe with fish broth) in extensive field studies of bacterial abundance, distribution and species composition. Fischer sought to test the hypothesis that bacteria played a major role in nutrient cycles in the ocean as they were already known to do on land. His culture-dependent results did not permit a quantitative test, but being impressed by the ubiquitous distribution of bacteria and their ability to degrade organic matter in the laboratory he concluded: when we learn that bacteria are preseJlt in ocean waters even in considerable depth, and when we observe that they can readily develop on dead animal and plant material .... then we can not doubt that these bacteria are as important causes of decomposition in the ocean as bacteria are o~I land ... (Fischet, 1894; translated from German by C. A. Painter).
As we shall see later, Fischer's intuition was correct. Generations of microbial oceanographers that followed have sought to test Fischer's hypothesis but methods to quantify in situ microbial activities were still lacking for another 80 years! Viable count remained the method of enumeration. The counts being low, typically around 10~ ml ', the resulting and persistent perception was that such sparse populations of bacteria could not be important in the ocean's ecological and biogeochemical dynamics. Since a low bacterial abundance was the basis of the persistent paradigm of the insignificance of the ecosystem role of bacteria it is pertinent to ask why a better method to count bacteria was not developed for so long? Indeed, several direct count methods (e.g. filter-concentration method of Cholodny, 1929) were known to yield two to three orders of magnitude higher counts than the viable counts. True, these methods were cumbersome and impractical for processing large numbers of field samples, but little effort was made to improve them. ZoBell (1946) observed: direct counts are attended by many technical difficulties and have limitations z~,hich restrict their usqhdness. A t best, direct counts y, ive data which only supplement and aid i~l the interpretation of results obtained by cultural proced u res.
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Also, it was thought that direct counts mostly represented dead and inactive bacteria that were not useful in considering the role of bacteria in ocean processes. According to ZoBell (1946):
The siguffficance ~f cou~lting the lmmber of microorganisms in a given enviromueut is often over-estimated. The populatknl is merely an expression qf the dynamic balance between the rate off production and the rate of destrltction ~f microolxanisms. The appraising of the roh' o[ microotNanisms in chemical, ~eological and biological conditions, the rate of reproduction and activity o{ the microotxanisms is a more important consideration than is the mHuber q[ micro-
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ZoBell was absolutely right about the need to measure in situ microbial activities. But it seems that obtaining reliable direct counts was not a priority, and this lack of priority; rather than the lack of flat filters and fancy fluorescence microscopes, is w h y we did not have a good method until 1977. Thinking about the relationship between technology and methods in marine microbiology Grieg Steward once told me, we were able to reliably count bacteria in seawater on Earth eight years ~TflerMan had landed on the Moon! Did the lack of suitable methods prevent the integration of microbial processes into oceanography? Reflecting on this point, Haldane Gee (1932) threw up his proverbial hands:
... the bacteriolo~,qst caJl not yet avail himself with profit of an oceam~raphic cruise. The field of mariHe microolxanisms has m~t been sufficieJ~t]y explored to /ust!fy the hope thai laborious sample aud culture work at sen could be expected to enlalxe the existing in.&rmation sufficiently. Moreover, the technical difficulties ... deter the conscientious investigator from such atl undertakills¢. It is admitted that volumillous data could be gathered by such work oil board ship. Whether the Sll~]/~osed bacterhT[ COUlltS so recorded woll/d be Off s@llJficallce, ]lowez~el', is a~other matter. (Although marine microbiology has n o w advanced to become an integral part of oceanography, there is still some validity to Gee's admonition against uncritically collecting 'voluminous data" in oceanographic studies.) It seems that for decades, until John Hobble (Hobbie et al., 1977) s h o w e d high bacterial abundance, microbial oceanographers were in an unenviable limbo. They could not integrate into oceanography, since oceanographers had no need for their CFUs or the physiology of their isolates. On the other hand, their findings were of little interest to classical microbiologists (except for their interest in some intriguing isolates from the sea,
e ,, Photobacteria). It is generally agreed that the renaissance in microbial o c e a n o g r a p h y began with the publication of L. R. Pomeroy's 1974 paper (Pomeroy, 1974), which then stimulated a blossoming of m e t h o d d e v e l o p m e n t to test his ideas. However, the ferment for change was derived from the 1960s which saw quite remarkable activity in method development. Hardly any of the methods has survived but their influence on future d e v e l o p m e n t of ideas and methods has been profound. So the 1960s were exciting not only for
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social activism but also for methods in marine microbiology, and curiously the two may have been connected. It was an era of keen awareness and activism concerning environmental issues. The lakes were dying, the oceans were polluted and the air was dirty. Environmental activism forced the recognition that too little was known about marine ecosystems to prevent or rectify ecosystem deterioration. Prevention would be preferable but it required the ability to predict the consequences of potential ecosystem insults (a goal yet to be achieved, and it is being discussed in relation to the modern pollutants, the greenhouse gases). How much toxic heavy metals could we dump into a bay without sickening the biota? Will DDT, PCBs and radionuclides entering the ocean contaminate the fish we eat? Are we stressing the marine ecosystems such that, like the lakes before, they too will 'die' of poisons, eutrophication and loss of biotic diversity? Answers to such questions required quantitative studies of the behavior of intact ecosystem, or at least in situ biotic activities, e.g. food web transformation and transfer of marine pollutants. The culture technique was not enough, and direct, quantitative ecosystem analysis was required. This was a tall order. And there was an urgency to find answers. This need for answers to sharply defined questions provided, for the first time, a specific agenda and resources which helped focus the energies of microbial oceanographers on culture-independent methods (and in the process expanded the field). A number of methods were developed to measure biomass, activity as well as the 'health' of marine microbiota and marine ecosystems. These methods were able to flourish in parallel with culture-based analyses of species diversity (e.g. numerical taxonomy) and its response to ecosystem stress. Phytoplankton methods were already relatively advanced: chlorophyll a for biomass, microscopy for species diversity and the ~4C method for primary productivity. So, stress response of phytoplankton communities could be quantified in terms of these parameters. Bacteria still could not (even) be counted; but could we measure their in situ activity and 'health' without having to count them? Deriving inspiration from the primary productivity method, a method was developed using the uptake of ~C labeled organic substrates as indicator of microbial heterotrophic activity (Parsons and Strickland, 1962). Wright and Hobble (1965) extended it into a kinetic approach to measure in situ substrate turnover rate and the V...... ('heterotrophic potential'). A radio-respirometry method (Hobbie and Crawford, 1969) was developed to measure ~4C substrate respiration. These methods ushered in the 'radiotracer era' in microbial oceanography, promising insights into iH situ bacterial metabolism, the role of bacteria in organic matter turnover and how pollutants affected these ecosystem roles of bacteria. While radiotracer method strongly influenced how we perceived microbial dynamics (below), it was not practical for quantifying carbon flux. DOM flux into bacteria involves many substrates and a piecemeal approach to individually measure uptake of all substrates was simply not practical. Also, the in situ substrate concentrations needed to be known to compute mass flux from tracer uptake of all potential substrates, but this was technically infeasible.
Despite these limitations, the method was applicable to measuring the turnover rates of specific sugars and amino acids and it was extensively used in oceanographic studies. The results were unexpected and exciting. The substrate pool turnover rates were very fast, much faster than expected from the presumed sparse and inactive bacterial assemblages. For instance, turnover times for sugars and amino acids in coastal waters were typically measured in hours! Even though the turnover rates could not be converted to mass flux, and we had no clue of bacterial abundance either, the rapid substrate turnover gave a feeling that DOM was dynamic, and that heterotrophic bacteria were quite active in mediating organic matter fluxes. This qualitative sense of dynamism of microheterotrophic processes provided the energy and set the stage for the revolutionary conceptual and technical changes which occurred in the 1970s and 1980s. Further, size-fractionation of tracer-labeled natural bacterial assemblages led to a second fundamental change in thinking, that most heterotrophic activity was due to those bacteria which were filterable through 1 t~m, even 0.6 ~m~, Nuclepore filters. This was contrary to the long-standing belief (since Waksman (1933) and ZoBell (1943)) that most bacteria were particle-associated and that bacterial activity was mainly restricted to particle surfaces. This issue has fundamental implications for bacterial ecology and organic matter cycling. (However, now it is being suggested that pelagic bacteria may not be completely unattached, since they may interact with DOM gel matrix or the cell surface mucus layers of other microbes (Azam, 1998).) The emphasis on culture-independent methods to study natural microbial assemblages also led to a flurry of biochemical methods for measuring bacterial (and total microbial) biomass and the physiological state of natural assemblages of marine microbes: ATP for total microbial biomass; lipopolysaccharide and muramic for bacterial biomass; electron transport system (ETS) activity as a proxy for microbial respiration; energy charge as a measure of the physiological state of the assemblage (for instance in response to ecosystem stress). Culture-independent methods must deal with the great challenge to measure a specific parameter of specific organisms which occur admixed with diverse assemblages of non-target organisms as well as detritus. As such, they did well, providing very useful constraints on microbial biomass and iH situ activities, and were extensively used in field oceanography and in studying ecosystem stress. These methods occupied center-stage in biological oceanography for a decade, and discussions on issues ranging from their conceptual bases to minute operational details elicited passionate debates during many a workshop and conference, it is remarkable, ahnost eerie, that they have disappeared from the scene as if they had never existed. Did they so fail? Did they simply go out of fashion? Or, did they give way to newer, more powerful and more incisive methods to unravel the ecosystem roles of marine microbes? I think that a combination of each caused their 'demise'. These methods can still be useful for specific questions but are rarely used because of the perception that they do not 'work'. Microbial respiration was another culture-independent method to assess the metabolism of natural assemblages in the 1960s and 1970s
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(Pomeroy, 1974; Williams, 1981). The method has had a 'boutique' status, a specialized method used by few. Yet, it is among the most powerful methods available to microbial oceanographers. Respiration of sizefractionated marine assemblages probably shaped Pomeroy's ideas on the significance of microbes as dominating pelagic metabolism. Williams (1981) used it to develop a quantitative view of the microbial oceanographic processes. Respiration provides important constraints on bacterial growth efficiency (del Giorgio and Cole, 1998). Method refinement to measure respiration of natural bacterial or protozoan populations would be a significant contribution to understanding microbial carbon cycling in the ocean. Unlike other methods of the 1960s some incarnation of respiration method is likely to play an important role in microbial oceanography in the future. The epifluorescence microscopy method (Hobbie et al., 1977) dramatically changed our view of the ocean's ecosystem. The method was developed to count bacteria, and the results were dramatic enough. Bacterial biomass was huge, three orders of magnitude greater than determined on the basis of viable counts. The 'bonus' discovery of abundant Synechococcus and protozoa increased the impact of the method. One might once again have dismissed the high bacterial abundance by arguing that most bacteria were dormant or dead and hence devoid of an ecosystem role; indeed, there was no new evidence to the contrary. But a different mind-set prevailed, and the direct counts were accepted at face value. Perhaps this was because of the sense of dynamism of microbial processes borne out of tracer studies, discussed above; or perhaps the acridine-orange-stained brightly fluorescent 'starry night' images of bacteria seemed so compelling and 'alive'. The sense of microbial dynamism was further reinforced by the next major discovery, made with the help of two new methods to measure bacterial production. As a backdrop, I should point out that prior to 1979 we had absolutely no idea how fast bacteria grew in the sea, whether they doubled once every hour or once every year! Did they grow fast enough to be a significant conduit for the flow of photosynthetically fixed carbon? The general sense was that bacterial production and carbon demand were so low that they would be responsible for utilizing only a trivial fraction of primary production. Metazoan grazers would then be responsible for much of the 'action' in energy and carbon flow. Was the role of bacteria really trivial in the structure and function of marine ecosystems? As already discussed, the piecemeal tracer turnover studies were inadequate for measuring the total organic matter flux into bacteria. Bacterial carbon production plus respiration could serve as a cumulative measure of total carbon flux into bacteria. Was this cumulative flux a trivial or a significant fraction of primary production? it is interesting to contemplate that on the answer to this question may have hinged the future of microbial oceanography! In a seminal paper published in 1979 Hagstrom and his collaborators (Hagstrom et al., 1979) reported an elegant method, the frequency of dividing cells (FDC) method, to measure bacterial production. Their results showed that bacterial production would have required on the
order of one-fourth of tile local c o n t e m p o r a n e o u s primary production. Karl (1979) published the ~'H adenine incorporation m e t h o d and Fuhrman and Azam (1980) published the ~H-thymidine incorporation method. The use of the ~H-thymidine incorporation m e t h o d in a mesocosm study showed that bacteria were responsible for the utilization of about one-half of the c o n t e m p o r a n e o u s primary productivity. Respiration in the bacterial size-fraction in the same mesocosms (Williams, 1981) showed that bacteria were responsible for roughly one-half of it. So, bacteria were major players in carbon and energy fluxes! The :H thymidine m e t h o d become popular and was widely used. But it has also been widely criticized. There are uncertainties of the conversion factor and several other technical issues some of which have been resolved and some have been forgotten or else accepted as a level of uncertainty acceptable in biological oceanography. Leucine incorporation m e t h o d (Kirchman et al., 1985; SimoFt and Azam, 1989), which more directly measures bacterial carbon production, via protein synthesis rate, later confirmed the conclusion that bacteria were major players in the flux of photosynthetically produced organic matter. These studies also established that bacterial populations were dynamic and created a fresh round of m e t h o d d e v e l o p m e n t to identify and quantify the sources of bacterial mortality. A method using H labeled bacteria lead to the still-current emphasis on the flow of carbon via the pathway: DOM --~ Bacteria -+ protozoa -+ metazoa, as well as the identification of bypass routes for high efficiency transfer (e.g. DOM -+ Bacteria ~ larvaceans ~ Fish) (Hollibaugh et al., 1980; King et aI., 1980). (The role of bacteriophages as significant killers of bacteria came a decade later; see below.) This view of the ecosystem role of bacteria finally consolidated the integration of marine microbiology with oceanography. No~.~; bacteria were demonstrably inw)Ived in a quantitatively significant way in processes central to biological oceanography, marine chemistry (DOM cycling), food web transfer of pollutants, and even fisheries and air-sea exchange of carbon dioxide. All these methods, all these discoveries - - starting from acridine orange direct counts to the almost current picture of the microbial food web - - all this happened in a short span of three years, 1977-1980! The m o m e n t u m and euphoria in microbial o c e a n o g r a p h y due to these discoveries has only increased over the last two decades. Clearl,,; the advent of molecular tools in o c e a n o g r a p h y is one reason for it (discussed later). But, perhaps equally important has been the energizing effect of the shear expansion of the field, because o c e a n o g r a p h y teaching and research programs quickly recognized the importance of microbial oceanography. In the mid-1970s Angelo Carlucci, who taught marine microbiology at Scripps, began his course with a map of North America showing the names and locations of American and Canadian marine microbiologists. The markings were pretty sparse. With more scientists in the field, it became possible for m a n y more facets of microbial o c e a n o g r a p h y to be studied and critiqued, i think that we are still too few to cover the major aspects of microbial oceanography. This view may seem rash to m a n y who think that we are already over-saturated, i believe that just as thus far new discoveries have opened new opportunities the same trend will
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persist. As I suggest at the end of this chapter, despite impressive progress, most major questions remain unanswered. And they are extremely important if we are really serious about learning enough about the Earth system to conserve it. Many more microbial oceanographers will be needed. The next phase of discovery might be called the decade of microbial exploration of the sea. It required new tools and techniques never before used in microbial oceanography. I will give two examples which caused fundamental change in oceanography. The development of ideas on microbial food web had focused attention on 'the small' in the sea. S. W. Chisholm (Chisholm et al., 1988) took the flow-cytometer to sea, to explore for phototrophic picoplankton. Her discovery of highly abundant Prochlorococcus marinus in the sea is one of the greatest discoveries in biological oceanography. It is remarkable that this picoplankter, which is apparently the most abundant known organism on Earth, had eluded detection for a hundred years of ocean observation. It also underscores the importance of direct observation of organisms in their environment. Further, the discovery of Prochlorococcus and Synechococcus showed that earlier methods had completely missed the major primary producers in vast oligotrophic areas of the world ocean. The final example of ocean exploration for the 'small' is the discovery, by electron microscopy, of highly abundant (10~-10~ml 1) viruses in the pelagic ocean, independently by two research groups (Bergh et al., 1989; Proctor and Fuhrman, 1990). Actually, there was indication of abundant viruses a decade earlier (Torrella and Morita, 1979) but the method was not optimized. New methods were rapidly developed to measure phage-induced mortality of bacteria and phage turnover. Impressive progress has since been made in understanding the biology and ecology of marine phage by the use of molecular methods and the development of new methods (Jiang and Paul, 1997; Fuhrman, 1999; Steward and Azam, 2000; Steward in press; Wommack and Colwell, 2000). The complete genome sequence of a -40 kbp marine Roseophage has recently been determined (Rohwer et al., 2000) to help infer its ecology. So, the progress has been rapid, partly due to the availability of molecular tools. One of most influential techniques in microbial oceanography is HansGeorg Hoppe's method for measuring bacterial ectohydrolase activities in intact planktonic assemblages (Hoppe, 1983). It has provided important insights and constraints on how the organic matter in particles and polymers become available for uptake by bacteria. In a broader sense, this method has contributed to the development of the interface between microbial oceanography and marine biogeochemistry. Its development as an individual cell technique is underway and should provide important bases for in situ interrogations at the single cell level. The development of a reliable and sensitive method for measuring protozoan grazing on bacteria and algae has been a challenge, judging from the large number of methods which have been proposed since 1980 (size-fractionation, dilution technique, fluorescently labeled prey, genetically labeled prey, differential metabolic inhibition, enzyme assays). While the current methods are able to provide useful constraints on
grazing rates the 'perfect' method is yet to be devised. This is an important future goal since protists play such a pivotal role in the flow of material and energy through the food web. The current emphasis on culture-independent microbial oceanography finds its most intensive expression in 16SrRNA-sequence-based phylogenetic analysis of marine prokaryotic communities. PCR amplification, cloning/sequencing, DGGE (denaturing gradient gel electrophoresis) and TRFLP enable analysis of phylogeny and species richness. The methodology enables one to study variations in species composition and richness in different environments or under different experimentally imposed conditions. The discovery of abundant as-yet-uncultured Archaea in the sea (DeLong, 1992; Fuhrman et al., 1992) is one example of the tremendous power of the methodology. Individual cell multiple interrogations in natural microbial assemblages promise to relate phylogeny with in situ physiology (Ouverny and Fuhrman, 1999; Cottrell and Kirchman, 2000), a most exciting prospect indeed. However, this methodology is not yet well established and should be a future priority. In fact, most of the molecular methods discussed here have limitation (e.g. interpretation of PCR and DGGE results of complex assemblages) but given the pace of progress in molecular biology it is reasonable to assume that methods will improve or better methods will rapidly supplant them. Another effect of molecular methods on oceanography has been to increase communication between microbial and other biological oceanographers through convergence of disciplinary methodologies. Everybody is running gels and doing PCR so molecular methods serve as an icebreaker. The power of the molecular methods is enormous and increasing. Genomic analysis of marine microbes promises to further revolutionize microbial oceanography. As I write, the complete genome sequence of Vibrio cholerae appeared today in Nature, and it should provide insights into the ecology of this estuarine and marine bacterium. Methodology is on the horizon to determine the genome sequence of as-yet-uncultured bacteria and viruses. Global gene and protein expression methodology should enable the response of bacteria to specific environmental interactions and stresses. Surely, the molecular and genomic approaches will revolutionize microbial oceanography. But we should not be seduced by molecular methods to the point where we define our research in terms of them. Future advances in microbial oceanography will also require developing new types of methods (below). Also, there is the issue of availability versus accessibility of methods. Funding for biological oceanography has traditionally been quite modest and does not support the 'kit based' molecular techniques. If my lab is an example of an average lab in microbial oceanography then there are significant monetary constraints on the accessibility of the available molecular methods. And I recognize that most microbial oceanography labs in the world are not as lucky as mine. We are creating a class system of sorts, the have and have-nots of microbial oceanography, the rich reviewers of manuscripts wondering why the author did not just go ahead and clone and sequence all those DGGE bands. There is no simple
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solution to the cost issue. The granting agencies must recognize that microbial oceanography n o w depends on costly molecular methods. If we are serious about the health of the oceans it will take serious money, the type of funding given for research on h u m a n health. As scientists we should do our share by better planning experiments and by minimizing our d e p e n d e n c e on 'kits'. Our colleagues, here and around the world, who are not well funded will have to make their own reagents as much as possible, but perhaps that will provide an aspect of training missing in the rich labs. More collaboration should also be helpful. What about the future? There are exciting conceptual challenges ahead which will require new methods. A powerful new emphasis in oceanography is to develop an integrative view of how biological forces pattern the flows of carbon and energy in ocean space and in time. While the current methods have shown that microbes greatly influences marine ecosystem structure and dynamics our knowledge is descriptive, and we cannot predict ecosystem behavior or its response to stress, e.g. 'global change'. In order to develop predictive capability we must u n d e r s t a n d the mechanistic bases of the interplay between the microbes and their environments. The microenvironment of the microbe is the big unknown; we know nothing about the chemical, physical or biological characteristics of the micrometer to millimeter scale worlds in which the microbes live and interact with other components of the environment to exert their influence. An exciting recent discovery is that the seawater environment of bacteria is a hydrogel structured by cross-linked polymers (Chin et al., 1998; Azam, 1998). Microbial ecology obviously occurs at the microscale, but microbial ecology has essentially been intractable until the advent of molecular biology. Intractable indeed since: we could not count or identify the microbes nor could we isolate but few; we had no methods to measure their growth and death rates; we could not find out 'what they were doing' in their environments, nor could we characterize their environments; we could not study their in situ physiology, behavior and interactions nor the consequences of these adaptive interactions for the structure and functioning of the ecosystem. We still do not k n o w the answers to most of these questions. So, w h y all this euphoria? It is because the new techniques, and those on the horizon, give cause for optimism that microbial ecology has become tractable for the first time. Individual-based ecology in an ecosystem context should be a focus for the future method development. Genomics and proteomics promise mechanistic and integrative bases of how the interplay between the expression of the microbes' adaptive strategies in their environment shapes the ecosystem at all scales of space, not just the microscale. Such knowledge will help produce a new type of ecosystem model, able to predict the biogeochemical behavior of the ocean u n d e r 'normal' and stressed conditions. Predictive models of ecosystem behavior should help provide sound solutions to societal issues such as marine pollution, toxic blooms, emerging diseases and coastal eutrophication. The methods in microbial oceanography have come of age, there is m u c h to do, and it is going to be terribly exciting! l0
References Azam, E (1998). Microbial control of oceanic carbon flux: The plot thickens. Science 280, 694-696. Bergh, f~., Borsheim, K. Y., Bratbak, G. and Heldal, M. (1989). High abundance of viruses found in aquatic environments. Nature 340, 467-468. Chin, W.-C., Orellana, M. V. and Verdugo, P. (1998). Spontaneous assembly of marine dissolved organic matter into polymer gels. Nature 391, 568-572. Chisholm, S. W., Olson, R. J., Zettler, E. R., Goericke, R., Waterburn, J. B. and Welschmeyer, N. A. (1988). A novel free-living prochlorophyte abundant ill the oceanic euphotic zone. Natmv 334, 340-343. Cottrell, M. T. and Kirchman, D. L. (2000). Natural assemblages of marine Proteobacteria and members of the Cytophaga-Flavobacter cluster consuming low- and high-molecular-weight dissolved organic matter. Appl. Environ. Microbiol. 66, 1692-1697. deI Giorgio, P. A. and Cole, J. J. (1998). Bacterial growth efficiency in natural aquatic systems. Amt. Rev. Ecol. Syst. 29, 503-541. DeLong, E. E (1992). Archaea in coastal marine environments. Proc. Natl. Acad. Sci. 89, 5685-5689. Fischer, B. (1894). Die bakterien des meeres nach den untersuchungen der planktonexpedition unter gleichzeitiger berucksichtingung einiger alterer und neuerer untersuchungen (Ergebnisse der plankton-expedition der Humboldtstiftung, 4, 1-83; Lepsius and Teicher) (Translated into English by C. A. Painter; edited by C. E. ZoBell and S. C. Rittenberg. Available at the library of Scripps Institution of Oceanography, University of California.) Fuhrman, J. A. (1999). Marine viruses: biogeochemical and ecok)gical effects. Nature 399, 541-548. Fuhrman, J. A. and Azam, E (1980). Bacterioplankton secondary production estimates for coastal waters of British Columbia, Antarctica and California. Appl. Envin~u. Microbiol. 39, 1085-1095. Fuhrman, J. A., McCallum, K. and Davis, A. A. (1992). Novel major archaebacterial group from marine plankton. Natmv 356, 148-149. Gee, H. (1932). Marine bacteriology - - scope and function. (Final report as bacteriologist, Scripps Institution of Oceanography, La Jolla, CA.) Hagstr6m, A., Larsson, U., H6rstedt, P. and Normark, S. (1979). Frequency of dividing cells, a new approach to the determination of bacterial growth rates in aquatic environments. Appl. Euviron. Microbiol. 37, 805-812. Hobble, J. E. and Crawford, C. C. (1969). Respiration corrections for bacterial uptake of dissolved organic compounds in natural waters. LimHo[. Oceano~r. 14, 528 532. Hobble, J. E., Dales, R. J. and Jasper, S. (1977). Use of nuclepore filters for counting bacteria by epifluorescence microscopy. Appl. Epr~qlxnt. Microbiol. 33, 1225-1228. Hollibaugh, J. T., Fuhrman, J. A. and Azal~n, E (1980). A technique to radioactively label natural assemblages of bacterioplankton for use in trophic studies. Limnol. Oceauogr. 19, 995-998. Hoppe, H.-G. (1983). Significance of exoenzymatic activities in the ecology of brackish water: measurements by means of methylumbelliferyl-substrates. Mar'. Ecol. Prog. Set., 11, 299-308. Jiang, S. C. and Paul, J. H. (1997). Significance of lysogeny in the marine environment: studies with isolates and a model of lysogenic phage production. Microb. Ecol. 35, 235-243. Karl, D. M. (1979). Measurement of microbial activity and growth in the ocean by rate of stable ribonucleic acid synthesis. Appl. EnviroH. Microbiol. 38, 850-860.
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King, K. R., Hollibaugh, J. T. and Azam, E (1980). Predator-prey interactions between the larvacean Oikopleura dioica and bacterioplankton in enclosed water columns. Mar. Biol. 56, 49-57. Kirchman, D., K'Nees, E. and Hodson, R. (t985). Leucine incorporation and its potential as a measure of protein synthesis by bacteria in natural aquatic systems. Appl. Environ. Microbiol. 49, 599-607. Ouverney, C. O. and Fuhrman, J. A. (1999). Combined microautoradiography-16S rRNA probe technique for the determination of radioisotope uptake by specific microbial cell types in situ. Appl. E~iviron. Miclvbiol. 65, 1746-1752. Parsons, T. R. and Strickland, J. D. H. (1962). On the production of particulate organic carbon by heterotrophic processes in sea water. Deep-Sea Res. 8, 211-222. Pomeroy, L. R. (1974). The ocean's food web, a changing paradigm. Bioscience 24, 499-504. Proctor, L. M. and Fuhrman, J. A. (1990). Viral mortality of marine bacteria and cyanobacteria. Nature 343, 60-62. Rohwer, E, Segall, A., Steward, G., Seguritan, V., Breitbart, M., Wolven, E and Azam, E (2000). The complete genomic sequence of the marine phage Roseophage S101 shares homology with non marine phages. Limnol. Ocemlogr. 45, 408-418. Simon, M. and Azam, E (1989). Protein content and protein synthesis rates of planktonic marine bacteria. Mar. Ecol. Prog. Ser. 51, 201-213. Steward, G. E (2001). Fingerprinting viral assemblages by pulsed field gel electrophoresis, this w~lume, 85-104. Steward, G. E and Azanq, E (2000). Analysis of marine viral assemblages, Microbial Biosystems: New frontiers. Bell, C. R., Brylinski, M. and Johnson-Green, P. eds, Proceedings of the 8th Int. Symposium on Microbial Ecology. Atlantic Canada Society for Microbial Ecology, 159 165. Torella, E and Morita, R. Y. (1979). Evidence by electron cnicrographs for a high incidence of bacteriophage particles in the waters of Yaquina Bay, Oregon: ecological and taxonomical implications. Appl. EHviroJz. Mierobiol. 37, 774-778. Waksman, S. A., Reuszer, H. W., Carey, C., Hotchkiss, M. and Renn, C. E. (1933). Studies on the biology and chemistry of the Gulf of Maine. III. Bacteriological investigation of the sea water and marine bottoms. Biol. Bull. 65, 83-205. Williams, P. J. L. (1981). Incorporation of micruheterotrophic processes into the classical paradigm of the plankton food web. Kieler Meeresforsch. Somh, r. 5, 1-28. Wommack, K. E. and Colwell, R. R. (2000). Virioplankton: viruses in aquatic ecosystems. Microbiol. Mol. Biol. Rev. 64, 69--114. Wright, R. T. and Hobble, J. E. (1965). The uptake of organic solutes in lake water. Limm~l. Oceanogr. 10, 22-28. ZoBell, C. E. (1943). The effects of solid surfaces upon bacterial activity. ]. Bacteriol. 46, 39-53. ZoBell, C. E. (1946). Marine Microbiology. A Monograph o~l Hydrobacteriology. Chronica Botanica Co.
12
2 Microbial Ecology at Sea: Sampling, Subsampling and Incubation Considerations D H Karl and JE Dore Department of Oceanography,School of Ocean and Earth Scienceand Technology,University of Hawaii, Honolulu, HI 96822, USA
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CONTENTS General introduction How to sample, and where When to sample, and how often Incubation experiments and rate determinations Ecosystem-level experiments The HOT program protocols: A case study Conclusions and prospectus
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GENERAL
INTRODUCTION
The marine environment is the largest contiguous habitat on Earth; however, it is far from being homogeneous. Relevant ecological time and space scales span more than nine orders of magnitude (Figure 2.1; Dickey, 1991), thereby contributing to the challenge of adequate and representative sampling of the marine environment. Many distinct marine ecosystems and their microbial assemblages have been identified and studied, ranging from ice-swept polar seas to deep-sea hydrothermal vents. The diversity of microbial habitats is even greater than already implied, especially considering the fact that microorganisms live in microenvironments that are defined on space scales of millimeters or less. As a result, the environments sensed by individual microbes may be quite different from the surrounding bulk fluid. Furthermore, many microbes live in truly protected habitats such as the enteric tracks of larger metazoan organisms; sampling these microbes will require fundamentally different methods than those used to target 'exposed' microbial assemblages. In fact, the spectrum of oceanic habitats and the diversity of the associated microbial assemblages is so extreme that any broad generalizations regarding sampling, subsampling and METt tODS IN MICROBIOLOGY, VOLUME 3(/ ISBN 0 12 521530-4
C o p y r i g h t © 2001 Academic Press Ltd All rights of reproduction in a n y form reserved
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Figure 2.1. Time and space scales of variability for physical and physiological processes that are relevant to marine microorganisms. The region beneath and to the right of the two arrows defines the time-space domain typically investigated by ship-based sampling programs. (From Dickey, 1991.)
measurement protocols must be carefully reviewed before application to a selected study site. Biogeochemical cycles of carbon (C), nitrogen (N) and phosphorus (P) in the sea are ultimately driven by solar energy and a continuous supply of growth nutrients. This results in a steep gradient in potential energy in the upper 0-100 m of the water column (the so-called euphotic zone, where net photoautotrophic fixation of carbon dioxide occurs), and a vertical segregation between net autotrophic and net heterotrophic microbial processes. Significant latitudinal variations in solar radiation, as well as vertical and coastal to open ocean horizontal gradients in nutrient concentrations are also present. Furthermore, within a selected habitat there are potentially significant did, intra- and inter-seasonal and interannual variations in microbial processes, and it now appears almost certain that decade-scale and longer climate forcing of the marine environment impacts the resident microbial communities (Tont, 1976; Venrick ef al., 1987; Karl, 1999). Microbiologists now recognize three major lines of evolution: Bacteria, Archaea and Eucarya (Woese, 1994). In the sea, these three domains have overlapping size spectra, physiological characteristics, metabolic strategies and ecological function. Consequently it is difficult to separate these groups except by use of novel molecular biological techniques that are only now being introduced into the field of microbiological oceanography.
14
Microorganisms, especially Bacteria and Archaea are ubiquitous in the marine environment and are truly the 'unseen majority': it has recently been estimated that there are more than 10-~L~microbes in the world ocean (Whitman et al., 1998). In addition to this sheer number, global ocean microbial biomass is also substantial and accounts for 0.6-1.9 x 10" g C (Karl and Dobbs, 1998). Approximately 75% of the total microbial biomass occurs in open ocean habitats with roughly half of that biomass distributed in the upper 0-100 m of the water column and the remainder in the deeper portions (>100 m) of the sea. With an average ocean depth of about 4000 m, this means that the concentration of microorganisms decreases substantially with increasing water depth. Although microbiologists have applied laboratory-based pure culture techniques to marine isolates for over 100 years, we are still lacking a comprehensive view of the ecology of microorganisms in the sea. The subliminal fear that the laboratory-based models were fundamentally different from the native populations now seems likely (Giovannoni et al., 1990) It was not until 1988 that the most abundant photoautotroph in the sea, Prochlorococcus marimts, was discovered and isolated (Chisholm et al., 1988, 1992). Even more recently, abundant marine planktonic Archaea have been observed (Fuhrman et al., 1992; DeLong eta/., 1994), but not yet cultured. DeLong et al. (1999) and Karner et al. (2000) have reported that the Archaea:Bacteria ratio approaches unity in deep waters (>500 m) of the north Pacific Ocean thereby documenting a large biomass of Archaea that until a few years ago were not even suspected to be present in 'normal' marine habitats. The most abundant planktonic bacterial and archaeal species in the sea have not yet been isolated, so their physiological characteristics and, therefore, ecological niches remain largely unknown. Why sample at all? There are at least two basic objectives in marine microbiology: (1) to isolate specific microorganisms or genotypes for subsequent study; and (2) to provide quantitative information on the distribution, abundance or metabolic activities of the resident microbial assemblages. The three most fundamental microbiological properties of a given ecosystem, community structure, total standing stock of living microorganisms (also known as total microbial biomass) and rates of metabolism or growth, are still far from routine measurements. These are the master variables in the sea of microbes. Consequently if one is interested in 'marine microorganisms' it is clear that the research question or hypothesis under investigation will dictate the types of samples that are collected, the sampling frequency in time and in space, and the selection of methodologies that are to be employed. This chapter will provide a few general guidelines on sampling, subsampling and other relevant field experimental design criteria.
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Principle Sampling is one of the most important, but often overlooked, aspects of oceanography. Because of the ease with which seawater or sediment is 15
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obtained, it is tacitly assumed that sampling is a straightforward and simple procedure. However, in our view, the task of obtaining an intact, representative sample of the marine habitat under investigation is the most significant challenge in microbiological oceanography. It is now well established that the pelagic realm of the world's ocean consists of readily identifiable habitats or biogeographic provinces (McGowan, 1974). These regions coincide with major hydrographical features, and can even be surveyed from space using color-sensing satellites (Platt and Sathyendranath, 1999). Accurate and precise descriptions of spatial and temporal patterns of microorganisms in the sea are fundamental parameters for marine microbial ecology. These objectives demand a rigorous and well-designed sampling program and appropriate methods of sample collection. A sample is intended to be just that, a representative subset of the population under investigation. Questions of time and space scale of variability of the population or habitat 'unit', sampling and measurement accuracy and precision and other relevant issues need to be considered before the experiment begins, perhaps using a 'pilot study' approach (Andrew and Mapstone, 1987). Some microbes of interest are large enough to be captured in nets (e.g. colonial or aggregated microorganisms like Trichodesmium or Rhizosolenia, and microorganisms associated with zooplankton) or in particle interceptor traps (e.g. microbes associated with rapidly sinking particulate matter). If plankton nets are used, it is up to the investigator to decide whether to tow the net at a single reference depth or over a specified depth range (i.e. a horizontal tow sampling design) or obliquely through a pre-selected depth stratum. For quantitative estimates, it is also crucial to measure the volume of water passed through the net. The larger the target organisms, in general, the fewer the number per unit water volume, and the more heterogeneous the distribution. Nets that can be opened and closed on command are preferred, and multiple opening and closing plankton samplers are ideal (B6, 1962). However, most microorganisms are unevenly dispersed in the bulk fluid habitat, and are collected using any one of a variety of water sampling devices. When designing an ecologically-based field program, care must also be given to the uncompromised collections of complementary data including dissolved substrates, dissolved gases, particulate matter and other parameters. In addition to the availability of numerous sampling devices, some of which will be described below, there is also a variety of potential sampling platforms, including boats and research vessels, towed vehicles and towed undulating vehicles, submersibles, remotely operated vehicles, autonomous underwater vehicles, moorings, drifters and Earth-orbiting satellites. Each platform has its own unique capabilities and limitations. Integrated measurement systems including multiple sampling platforms and fast response chemical and microbiological sensors are likely to emerge as the method of choice in future investigations of microbial processes in the sea (Dickey, 1991). The critical importance of adequate sampling of the ocean environment can be traced back to the early nineteenth century, when serious misconceptions about the deep-sea environment were presented. A respected 16
naturalist, Edward Forbes (1815-1854), claimed to have proven that below approximately 600 m in the open sea there was a 'probable zero of life' zone (Schlee, 1973). From an observed decrease in the number of animal species with increasing water depth, he concluded that the ocean was 'azoic,' or devoid of all life at great depths. His azoic zone theory was not refuted until the 1860s when a deep-sea cable from 2000 m was raised for repairs and revealed the presence of encrusting organisms (Gross, 1972). Clearly there had been a serious 'sampling problem.' As mentioned above, the ocean is not a single homogeneous ecosystem, so careful consideration must be given to sampling frequency (in time and space) and location. The scale of sampling relative to the scales of variability is important if one plans to extrapolate results to the ecosystem level (Levin, 1992). In most field studies, usually due to practical considerations, the actual number of total samples collected is regrettably small, and it is often impossible to obtain truly replicate samples. If statistical methods are employed, it must be assumed that the microbial populations follow a known probability distribution (e.g. Poisson, negative binomial or log-normal). However, microoganisms generally exist in localized patches and are rarely, if ever, found in random or uniform distributions over the spatial scales used in most ecological investigations (Karl, 1982). The investigator should be aware of at least three separate areas where variability can be introduced into field measurements: replication at tile level of sampling (i.e. multiple water samples collected from a common depth); replication at the level of subsampling (multiple subsamples from a single sample); and analytical replication (i.e. multiple analyses of a single sample extract). Because of the heterogeneous distribution of microbial communities in nature, and problems that are inherent in the collection of particulate matter from aquatic environments, variance between sampling bottles is generally the largest source of error. Therefore, replication is most meaningful when performed at the highest level, i.e. multiple samples of water from a given environment (Kirchman et al., 1982). It has also been demonstrated that the overall variance and the precision with which the sample variance can be estimated are functions of the procedure used to subsample the initial sample collection (Venrick, 1971). The need for a device that is capable of aseptically collecting a sample of seawater was recognized more than 100 years ago as the field of marine microbiology emerged as a subdiscipline of oceanography. ZoBell (1941) provides a thorough historical account of the significant events during the period 1892-1940, beginning with the pioneering work of H. Russell and W. Johnston. Over the years, a variety of aseptic water samplers have been devised, deployed and re-evaluated. There are two basic approaches used in their design: (1) a device using a capillary tube inlet that is deployed in a sealed, sterile configuration, opened at depth and recovered without closure and (2) the use of a mechanical device for the removal and subsequent replacement of a stopper or similar closure mechanism (Lewis et al., 1963). In 1941, ZoBell introduced the Johnson-ZoBell (J-Z) bacteriological sampler (which of course could also be used to sample Archaea and 17
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Eucarya). It consisted of a sterile, evacuated glass bottle, fixed stopper and glass and rubber tubing leading to a terminal sealed glass tube. The entire apparatus could be autoclaved at sea, then mated to a brass frame and attached to a hydrowire. A brass weight, also called a messenger, designed to travel down the same hydrowire on command, mechanically activated a lever that broke the glass tube; a water sample was aspirated into the sterile, evacuated bottle. The glass bottle could be replaced by an evacuated, compressible rubber bulb for greater depth capability (the glass bottles began to fail at about 200 m and by 600 m all bottles were crushed by the ambient hydrostatic pressure; ZoBell, 1941). A version of this modified J-Z sampler was also designed as a piggy-back microbiological sampler for use with the metallic Nansen bottle (Sieburth et al., 1963) which was the most commonly used water collection device for many years (pre-1965). Most of the early water samplers had integral components that were constructed from alloys of copper, nickel, tin, zinc or lead, even though the bactericidal effects of certain metals was well known (Drew, 1914). The rubber bulbs used in the modified J-Z and piggy-back samplers were also shown to be toxic to certain microbes. A substantially different sampler designed for the aseptic collection of seawater was introduced by S. Niskin and was termed the butterfly baggie sampler (Niskin, 1962). This bellows-type water sampler consisted of a spring-activated metal frame and a detachable 2 1 sealed, sterile, disposable polyethylene baggie. On command, the messenger activated a non-sterile knife blade which cut the end seal of the inlet and released the torsion springs opening the bellows. This action created a suction and collected a water sample from any target depth in the ocean. A mechanical, spring-loaded system then resealed the inlet tube at the completion of the sampling routine to prevent water exchange during sample recovery. One reported problem with this sampler was the leakage of dissolved organic matter from the plastic bags (Sieburth, 1979), so even though the sample is collected aseptically it may not be uncompromised for certain ecological measurements. Despite these developments, there was still concern expressed that these 'sterile' samplers might be compromised because of the requirement for in situ inlet tube activation by non-sterile procedures. Furthermore, the proximity of the intake of the sterile sampler to potentially contaminating non-sterile and metallic surfaces, including the hydrowire itself, raised suspicion regarding the reliability of these collections. For these reasons, Jannasch and Maddux (1967) developed a novel device consisting of a sterile syringe and glass sampling tube; the latter enclosed in a dialysis bag filled with sterile water. The apparatus is mounted on a movable arm that is attached to a frame and secured to the hydrowire (see Figures 2 and 3 in Jannasch and Maddux, 1967). A vane keeps the syringe oriented upcurrent to minimize potential contamination. When the messenger activates the sampler, the movable arm begins to fall away from the frame and its motion strips the protective dialysis bag away from the sterile glass sampling tube. At the same time, a cable attached to the syringe plunger begins to tighten and precisely when the arm is at its maximum distance from the frame assembly (approximately 75 cm) the seawater 18
sample is aspirated. Field tests of this sampler showed a greater reduction in the recovery of contaminating bacteria (identifiable bacteria that were deliberately 'painted' onto the hydrowire for these tests) than was observed with the paired deployment of either a modified J-Z sampler or a sterile Niskin baggie sampler (Jannasch and Maddux, 1967). Ironically, this relatively simple and effective aseptic water sampler was never used extensively in subsequent field programs. This might have been due to the emergent views of that time that the need for aseptic samples, when a relatively clean and uncontaminated one would suffice, was not necessary (Sieburth, 1979). Most of our conceptual views of the marine environment and, therefore, the basis for our sampling protocols focus on vertical profiles of oceanic parameters despite the fact that the marine environment is decidedly a 'horizontal' habitat (e.g. the horizontal-to-vertical scale of the North Pacific Ocean is >1000:1). In the open sea, this experimental protocol bisects a density-segregated water column with specific, readily identifiable layers called water masses. These components of the vertical profile vary considerably in their source and, most likely, in their chemical and microbiological properties. For example, a relative peak or m i n i m u m value for some selected parameter in a given vertical depth profile could be either an i~l sitlt or an advective feature. Clearly, an accurate resolution of these opposing mechanisms is desirable if not mandatory. In the North Pacific subtropical gyre at Sta. ALOHA (A Long-term Oligotrophic Habitat Assessment), the subeuphotic zone water mass at approximately 300 m (North Pacific Intermediate Water) is formed in the NW Pacific near Japan, but the deep water mass (>3000 m) has its origin in the Southern Ocean. Mixing between water masses occurs primarily at their boundaries so it is crucial to understand and consider this vertical structure w h e n designing a sampling program. If horizontal sampling is desired, one must be cognizant of the orientation of the plannned transect (zonal vs. meridional) and should anticipate changes in the water mass structure and positions as the source regions are approached. Because these water masses can be dated using transient chemical and radiochemical tracers, a well-designed transect can provide information on rates of change (e.g. O~ consumption, net nutrient uptake or regeneration, net bacterial production) in this 'upstream-downstream' sample design. The use of rosette-mounted water bottles and a CTD-based environmental sensing system provides for the real-time detection of water mass structure. In the open sea, there is a predictable vertical zonation of microorganisms including the following well-defined macro-habitats: (1) air-sea interface; (2) euphotic z o n e ; (3) mesopelagic zone; (4) abyss; (5) water-sediment interface; and (6) sediment column. The latter topics, including both the water-sediment interface and deeper subsurface sediments will not be discussed in this chapter. While there is ample evidence to conclude that marine sediments, in both coastal and abyssal habitats, support an elevated concentration of microorganisms relative to the overlying seawaters, detailed ecological studies of microbial processes in these important habitats are severely methods- (both sampling and analysis) limited. For these reasons we will focus on the water column. 19
_o o I,l,I ° J
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The air-sea interface, defined as the upper 150-1000 btm of the seawater, is a unique habitat characterized by high surface tension, high light (especially UV-B radiation), variable temperature, salinity, and turbulence. This specialized habitat also has generally elevated concentrations of dissolved organic matter, trace elements and microorganisms (Dietz et al., 1976; Sieburth et al., 1976; Carlson, 1982b; Williams et al., 1986). Depending on the assumptions used for the thickness of the sea surface microlayer, the enrichment factors (i.e. the concentration in the microlayer compared to submicrolayer surface water) can be 102-103, or greater. Bubble scavenging of surface-active organic matter and microorganisms is probably one important mechanism for sustaining these enrichments (Bezdek and Carlucci, 1974; Blanchard and Syzdek, 1982). There is no doubt that the surface microlayer habitat and, presumably, its microbial inhabitants, are fundamentally different from the underlying euphotic zone. Because the skin of the ocean is so important for heat, momentum and mass exchange, including gas fluxes, this under-studied habitat may be very important in issues related to global environmental change (GESAMP #59, 1995). The sea surface microlayer habitat is most likely composed of a series of overlapping zones that are difficult to sample quantitatively. Over the years a variety of instruments have been used, including: (1) the prism dip (Baier, 1972); (2) screen sampler (Garrett, 1965); (3) rotating ceramic drum (Harvey, 1966); (4) stainless steel tray (Hatcher and Parker, 1974); and (5) glass plate sampler (Harvey and Burzell, 1972). The efficacy of these methods has been evaluated in the laboratory (Hatcher and Parker, 1974; Van Vleet and Williams, 1980) as well as under field conditions (Carlson, 1982a). A mobile platform for studying the sea-surface film has also been described (Williams et al., 1982). The euphotic zone of the ocean is probably the most well-studied region with regard to microorganisms. Although vertically stratified, the seawater can be easily sampled and subsampled in time, using any one of a number of commercially available or homemade water collection devices. The water sampled with these devices includes dissolved constituents, viable planktonic microorganisms and some fraction of the total non-living particulate matter pool. There are at least two fundamentally distinct classes of particles in the sea: (1) particles that sink or rise and (2) particles that are approximately neutrally buoyant in the water column. Only those that are nearly neutrally buoyant can be sampled effectively with water bottles. The remainder of the particulate matter inventory must be collected using specialized devices such as sediment traps or large volume in situ pumps (Gardner, 1997). Depending upon the experimental objectives, the sample volume can range from <1 ml to >30 1; the former to sample discrete microenvironments (DiMeo et al., 1999), and the latter for routine sampling of dissolved and particulate matter. The most commonly deployed water sampler in contemporary microbiological oceanography is the Niskin ®bottle (or its equivalent) which in its simplest configuration is a polyvinyl chloride (PVC) cylinder with end caps secured by an internal elastic cord or spring. This non-sterile sampler is usually deployed open, by attachment to the 10
hydrowire, lowered to the target depth, then mechanically triggered to close by a messenger. Variations on this thellae include more elaborate bottle designs that can pass through the sea surface microlayer prior to opening at depth by a pressure-activated switch, to external closure mechanism bottles designed to reduce the potential for chemical contamination. Of course, like any other sampling device, the 'devil is in the details,' and so it is with water sampling bottles, especially with regard to the aspect ratio and inherent flushing characteristics (Weiss, 1971). Concerns have also been expressed about the potential incomplete recovery of large particles due to the positioning of the drainage spigots (Gardner, 1977). To the extent that microorganisms are unevenly distributed between freeliving and attached forms, this mechanical sorting and subsampling selection against large particles may be a significant source of sampling error. Most modern oceanographic investigations deploy a carousel of bottles attached to a large 1-2 m diameter circular frame referred to as a rosette. Typically, these 12-24 bottle packages are activated from the surface vessel using an electrically-controlled device on the rosette called the pylon. An essential requirement for this type of water sample collection is a hydrowire with electrical conductors and usually an environmental sensing device, termed a conductivity-temperature-depth (CTD) instrument, to provide real-time information on water depth and other habitat characteristics. Modern CTD devices provide the option for including additional underwater environmental sensors for relevant ecological parameters such as sunlight, light absorption and scattering, fluorescence and dissolved oxygen. These real-time data are invaluable for positioning the water bottles in zones of greatest potential interest (e.g. fluorescence maximum, particle maximum, O, minimum). Rosette-assisted sampling of the water column also provides for multiple water bottle sampling at a single depth, as needed for statistical evaluation or for large volume demands. Both hydrowire and rosette-mounted PVC water bottles can be thoroughly cleaned wifll 1 M hydrochloric acid and rinsed with distilled water prior to use. To our knowledge, there are no readily available procedures for the truly aseptic collection of large volume (>5 1) samples using rosette-assisted protocols. Other ingenious devices such as the tidalpowered (Hayes et al., 1980) and osmotic pressure-driven (Jannasch et al., 1994) water samplers can be used for unattended time-series collections. Sampling the marine environment at depths greater than approximately 2000m (the abyssopelagic and bathypelagic zones) requires special considerations and, depending upon the expedition objectives, sophisticated sampling gear. The successful isolation of deep-sea bacterial isolates that are obligately barophilic (Yayanos et al., 1979, 1981) has emphasized that pressure is an important determinant of microbiological zonation in the sea. Although some obligately barophilic bacteria may survive decompression, others may not. This implies that we may still have an incomplete understanding of microbial processes in the abyss. As Yayanos (1995) remarked, 'An ideal microbiological sample would remain in the dark, at the temperature and pressure of the deep sea, and mechanically and chemically undisturbed.' This is, unfortunately, generally not practical. While specialized pressure-retaining devices, including both 21
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water samplers and macroorganism traps, have been devised and deployed, the sample eventually needs to be decompressed for subsequent processing. To circumvent this problem, Jannasch et al. (1973, 1976) have developed a 1 1 pressure- and temperature-retaining deep-sea water sampler that, in connection with a transfer unit for the addition and withdrawal of 13 ml subsamples, can be used as an incubation vessel aboard ship or in the laboratory. A modified version of the deep-sea sampler can concentrate the water sample several hundred-fold, by in situ filtration (Jannasch and Wirsen, 1977). A high-pressure chemostat for field and laboratory-based studies has also been described (Wirsen and Molyneaux, 1999). However, these prototype devices are not commercially available and therefore are not generally employed. Recently, a sterile, rosettemounted high-pressure sampler capable of serial subsampling without decompression has been described (Bianchi et al., 1999) and employed to evaluate the pressure effects of microbial assemblages from the NW Mediterranean Sea (Tholosan et al., 1999).
O~l,e~l,~l,O W H E N T O SAMPLE, A N D H O W O F T E N
Principle The ocean is an ephemeral ecosystem, both in terms of the physical habitat and the microbial assemblages. This is especially true when one considers the in situ generation times of most bacteria (<1 day) and their potentially rapid response to perturbation. Even the duration and frequency of the E1 Nifio Southern Oscillation (ENSO) system, which itself exhibits a 3-5 year frequency of occurrence with well-documented effects on biogeochemical processes (Chavez et al., 1999), varies substantially on the decadal scale (Trenberth and Hoar, 1997). The impact of these lowerfrequency changes on ocean biogeochemistry remains largely unknown. The ENSO observing system in the equatorial Pacific Ocean, which includes moored and drifting buoys, a network of voluntary observing ship lines, and satellite surveys, is in place to observe the coupled physical-biogeochemical impacts of ENSO cycles. Furthermore, variation in time can produce coherent variation in space (Bennett and Denman, 1989); consequently separation of temporal and spatial variability of a given study region may be artificial (Jumars, 1993). Persistent coastal or equatorial upwelling, and the presence of quasistationary open ocean frontal regions such as the N Pacific Subtropical Front and the Polar Front, are fairly well studied. Less well documented are the sources of mesoscale (10-100 km) variability in the pelagic habitat. Direct evidence for the role of mesoscale eddies as an important control on nutrient delivery and sustained microbial primary and secondary productivity, especially in chronically starved, low nutrient open ocean ecosystems, has recently appeared (Falkowski et al., 1991; McGillicuddy et al., 1998; Oschlies and Garqon, 1998). These stochastic, pulsed processes disrupt the ecosystem steady-state and can lead to the selection of fundamentally distinct microbial assemblages. Detection of these short-lived 22
processes and integration of their effects into our conceptual models is only now starting to be achieved. There are at least two aspects of oceanic habitat variability that are relevant here. First, in order to ensure that a given sample is representative of the ecosystem under investigation, it may be necessary to obtain multiple samples in time. In tile open ocean there may be significant changes in heterotrophic bacterial processes that are coupled to the diel production of utilizable organic substrates by photoautotrophs. Likewise, in coastal habitats there may be a significant tidal influence on microbial processes that may be out of phase with diel periodicity. Probably all marine habitats, including tlle deep sea, experience intra- and inter-annual variations in the distribution, abundance and metabolic activities of the resident microbial assemblages. How many samples do we need to define this well-documented and fully expected temporal variability? There is, unfortunately, no simple answer to this question. Because seasonal changes in many marine ecosystems are substantial, the annual changes that integrate them from year to year are best evaluated over periods of decades to centuries. However, comprehensive longterm (>10 year) time-series observations of marine ecological or biogeochemical processes are rare. The >50 year Hardy continuous plankton record from the North Sea and North Atlantic and the >50 year plankton observations collected off Mexico and the southwest US coast as part of the California Cooperative Ocean Fisheries Investigation (CalCOFI) program are notable exceptions. Both of these studies focused, ultimately, on pelagic fisheries and neither contained a comprehensive study of the microbial assemblages or their controlling growth substrates. Relationships between upwelling intensity and plankton production was one physical link that emerged. In 1988, we began a systematic examination of microbial and biogeochemical processes in what was, at that time, considered to be a temporally stable habitat - - the North Pacific Subtropical Gyre. After the first decade of approximately monthly research cruises it was concluded that this sampling frequency was too coarse in time to fully resolve even the most important physical-biological interactions (Karl, 1999). The greater the number of time periods and space scales that are involved, the greater the measurement intensity to achieve even a basic understanding. As Stommel (1963) cautioned, 'Where so much is known, we dare not proceed blindly - - the risk of obtaining insignificant results is too great.' Undersampling is, unfortunately, a sobering fact of life in microbiological oceanography.
I N C U B A T I O N E X P E R I M E N T S A N D RATE DETERMINATIONS
Principle Many techniques currently employed in microbiological oceanography require incubation of a seawater sample for various periods of time. The 23
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underlying assumption of these methods is that the subsequent incubation conditions do not alter the in situ rates of metabolism or biosynthesis. This assumption is usually impossible to verify. Consequently it is imperative that the water sample is collected without chemical or environmental perturbation, and is subsequently incubated under conditions that duplicate those of the native habitat. Most microbiologists have a greater appreciation and concern for an aseptic sampling technique than they do for a chemically-clean sampling technique. Both are equally important but rarely enforced in field studies. For example, the most commonly used water sampling devices such as PVC water bottles are neither sterile nor necessarily free from contaminating materials. Furthermore, even if all viable microorganisms associated with a particular sampling device are killed, there is no assurance that specific cell biomarkers (e.g. lipopolysaccharide, nucleic acids) have been eliminated. Likewise, although an ethanol rinse might be recommended as a method to sterilize a sampling device or a subsampling instrument, it could grossly contaminate the sample with dissolved organic matter and otherwise preclude any reliable postsampling incubation procedures. Although an aseptic technique is probably not required for most routine field studies, to determine the presence or absence of a specific microorganism such a technique is imperative. An equally important concern, especially for post-collection incubation measurements, is attention to a clean sampling technique. Metal samplers, toxic closure components or other potentially detrimental materials should be avoided. Carpenter and Lively (1980) and Fitzwater et al. (1982) have warned of point source contamination by toxic trace metals during water sampling and subsampling procedures. This potential problem is especially acute when sampling surface waters of the open ocean where trace element concentrations are very low. Likewise, butyl rubber, latex rubber and neoprene tubing and o-rings should also be avoided because they have been shown to be very toxic to marine microorganisms (Price et al., 1986; Williams and Robertson, 1989). Only silicone materials appear to be acceptable and most chemically-clean samplers are not sterile. Depending upon the precise objectives of the particular study, one or both of these requirements can be ignored. However, while one should let common sense dictate, it is equally critical to know when and where point sources of microbial or chemical contamination are likely to occur. In order to ensure that the rates measured during the post-collection incubation procedure are representative of those occurring in nature, several precautions must be taken. First and foremost, the initial sample must be collected with great care so as to minimize chemical and microbiological contamination. Furthermore, exposure of viable microorganisms to environmental conditions that are substantially different from those at the collection site should be avoided so as to minimize any deleterious effects ranging from short-term transitions in metabolism to death. For selected habitats this will be virtually impossible, as described above for samples derived from deep-sea habitats. 24
A major limitation in microbial ecology is the lack of absolute standards or certified reference materials that can be measured along with the incubated samples. This places the entire burden for accuracy on the investigator, so the sampling and incubation design then become even more critical. In this regard, multiple complementary and even redundant assays should be performed, preferably using independent sample collections and incubation protocols, so as to constrain the specific ecological processes under investigation. The use of exogenous isotopic tracers has become a routine procedure for most field studies in marine microbiology. Often this is the only approach that is sensitive and specific enough to measure the sometimes low fluxes of carbon and other bioelements that occur in natural ecosystems. For example, the use of HC-bicarbonate as a tracer for carbon in marine photosynthesis (the so-called "C method; Steemann Nielsen (1956)) is probably the most widely used method in biological oceanography. Other exogenous isotope-based methods also exist for tracking heterotrophic microbial processes. The details of selected individual methods are discussed elsewhere, however there are several general considerations regarding the use of stable and radioactive isotope tracers in studies of microbial ecology that merit attention. These include: (1) the overall reliability of the added element (or compound) as a tracer, including an evaluation of the site of labeling, its uniqueness and stability during cellular metabolism and biosynthesis; (2) isotope discrimination factors; (3) the partitioning of the added tracer with existing exogenous and internal pools of identical atoms, molecules, or compounds and the importance of measuring the specific activity of the incorporated tracer; and (4) the design and implementation of experimental procedures and proper kinetic analysis of the resulting data. The use of isotopic tracers, especially radioisotopes, in ecological studies is often perceived as being straightforward and well documented. Consequently, a detailed working knowledge of the basic chemical, physical, statistical, and analytical principles on which these methods are founded is generally considered unnecessary. However, without such basic background information, it is possible to commit inadvertent errors in design or sample analysis that could result in gross misinterpretation of experimental data. In using commercially available isotopes, one relies to a large extent on the manufacturer's claims regarding the radiochemical, radioisotopic, and chemical purity of the product, tile position and pattern of labeling, and the measured specific radioactivity. Foreign chemicals are sometimes added deliberately to labeled compounds in order to improve the chemical stability (e.g. antioxidants) or radiochemical stability (e.g. radical scavengers) or as bactericidal agents. Impurities may also arise during the chemical, enzymatic or it~ vivo microbial synthesis of the specific compound or as a result of chemical hydrolysis (especially during long-term storage) and radiolysis. For example, both organic ~C (Williams et al., 1972; Smith and Homer, 1981) and trace metals, including Mn, Zn, Cu, Ni, Pb and Fe (Fitzwater et al., 1982), have been detected as contaminants in commercial preparations of [~4C]sodium bicarbonate 25
used routinely for primary production estimation. The presence of these contaminants may grossly affect the reliability of the tracer for measuring the rate of photosynthesis in aquatic environments. Another potential problem in tracer experiments results from the common use of ~H-labeled organic molecules as auxiliary labels for carbon. The primary advantage of the ~H-labeled compounds is their extremely high specific radioactivities, general availability and relatively low cost (per Bq). The former consideration is especially important for many ecological applications to avoid problems arising from organic nutrient perturbation resulting from the addition of the tracer (Azam and Holm-Hansen, 1973). The major disadvantage is the tendency of certain C-~H bonds to exhibit exchange reactions with H in the solvent (generally H~O). This exchange reaction may occur without any chemical change in the organic compound. Such labilization of tritium can occur due to chemical or enzymatic release during the intermediary metabolism of the tracer, during sample storage or during the extraction, purification and isolation of intermediate precursors or products. A very important but often overlooked principle in the use of isotope tracers in marine ecological studies is the evaluation of the specific activity (or atom % enrichment) of the added, incorporated or metabolized element, molecule or compound. The ideal tracer is one that can be added without perturbing the steady-state concentration of the ecosystem as a whole. The supplier is generally the source for specific labeling information; however, errors of up to 500% in the quoted specific activities of tritiated nucleosides from one supplier have been reported (Prescott, 1970). A detailed discussion of numerous potential sources of error in the calculation of specific activity has been presented by Monks et al. (1971). In ecological studies, an accurate assessment of the specific activity is further complicated by the dilution of the added tracer with exogenous pools present in the environment and by endogenous pools present in living microbial cells. Without a reliable measurement of the extent of dilution prior to incorporation, tracer uptake data by themselves are of limited use in quantitative microbial ecology. Furthermore, isotope specific activities may change over the course of the labeling period due to the combined effects of depletion (uptake) of the added tracer or isotope dilution by a constant regeneration of the exogenous pools (assuming steady-state conditions). In fact, NH4 + (Blackburn, 1979; Caperon et al., 1979) and HPO4~ (Harrison, 1983) regeneration rates have been estimated in environmental samples by measuring the extent of isotope dilution during short-term sample incubation periods. The final point of concern regards the theoretical bases and mathematical formulations required for the proper interpretation of data arising from the use of isotopes. This topic has been summarized and explicitly discussed by Smith and Homer (1981), who are of the opinion that ecologists in general and marine biologists in particular are largely ignorant of the vast body of literature available regarding the proper use of stable and radioactive isotopes. Smith and Homer (1981) present several multicompartment models and discuss the assumptions, restrictions and advantages of each approach. This type of rigorous kinetic treatment of tracer 26
data is only now becoming recognized as an essential component of the study of microbial processes in nature. A final consideration is the incubation itself; what is the best method for obtaining reliable rate estimates? Unfortunately there is no simple answer, but there are some recommendations to consider. First and foremost, the incubation conditions should match, to the extent possible, the sampled habitat. This simulation should mimic iJ1 situ light, temperature, pressure and all related chemical conditions. If a sample is collected from a lighted habitat, the incubation should also provide a similar flux of photons, even if the desired measurement is not directly coupled to sunlight. This is especially critical for measurements in ecosystems with a rapid turnover and, therefore, tight coupling between photoautotrophic and heterotrophic processes. If heterotrophic bacterial production is measured in the dark, it may underestimate the true in situ rate by depriving the heterotrophs of contemporaneously produced organic substrates. When in doubt, light vs. dark replicate treatments should be performed to assess the potential impact of this effect. In general, in situ incubation of samples is preferred to shipboard deck incubations. The in situ protocol ensures a match for temperature as well as light quality (wavelength) and quantity (Lohrenz et a/., 1992). Even when water samples are incubated under in situ or simulated (shipboard) in situ conditions there is a potential for metabolic perturbation during the initial sample collection process and subsequent pre-incubation handling. In order to circumvent the effects of light, temperature and pressure shock and in an attempt to obtain the most reliable estimation of the true in situ rate of primary production, Gundersen (1973) designed a clever device that he termed the in situ incubation sampler or ISIS (ISIS is also the Egyptian goddess of fertility!). The chemically-clean, non-metallic (but non-sterile) sampler is deployed on a hydrowire or synthetic rope to the depth of interest; multiple samplers can be deployed on a single wire. Each sampler has one opaque (PVC) and one transparent (polycarbonate) chamber. A sealed glass ampoule containing the HC-bicarbonate radioisotopic tracer is positioned in a special holder. When the sampler is activated by a messenger, the spring-loaded motion of the end-caps breaks the glass ampoule, thus inoculating the sample with the radioisotopic tracer and initiating the in situ incubation. The device remains in position for the duration of the pre-determined incubation period before it is recovered and processed. Dandonneau and Le Bouteiller (1992) have devised a 'clean,' ilk situ sample collection-incubation device which they termed 'let-go' because of its relatively simple deployment requirements. When natural populations of aquatic microorganisms are contained in glass bottles for periods of approximately 24 h, the composition of the population and rates of metabolism can change drastically and elicit the so-called 'bottle effect' (ZoBell and Anderson, 1936; Venrick et al., 1977). Bottle effects are totally unpredictable and vary considerably among taxa, location of sample and incubation conditions, in this regard, it would seem advantageous to keep incubations to a minimum duration that would still satisfy the prerequisites of the individual method (e.g. sensitivity in uptake, production or release of substance being measured, 27
intracellular radiotracer precursor equilibration). Even during relatively short-term incubations (1-3 h), there is the possibility of a nonlinear timecourse of metabolism as a result of confinement. One potential drawback with in situ experiments is the fact that they are generally end-point determinations and, therefore, lack information on changes that might occur during the incubation period. A recently devised sampler incubation device (SID) now provides the opportunity for an in situ, time-course incubation (Taylor and Doherty, 1990). Comparison of data collected with this method to standard end-point measurements have documented a serious, potential problem with prolonged end-point incubation experiments (Taylor and Doherty, 1990).
E C O S Y S T E M LEVEL E X P E R I M E N T S It is often desirable to deliberately manipulate the nutrient, trace element or other metabolic status of natural microbial communities in order to study their physiological and ecological responses over both the short (minutes to hours) and long term (days to weeks). However, as Strickland (1967) lamented, 'the open sea is too big; the laboratory beaker is too small.' Research using large plastic enclosures in the open sea has demonstrated the merits of this experimental design especially for the study of nutrient dynamics, organic matter production and recycling and the response of the microbial assemblages to short-term perturbations (Kuiper, 1977; Menzel and Case, 1977; Steele, 1979; Davies, 1984). The enclosures, though not meant to duplicate the habitat under investigation, are analogous to controlled experimental plots used in agricultural research (Menzel and Steele, 1978). In selecting a specific experimental design it is important that the volume selected is large enough to contain all components of the microbial food web including predators up to a few millimeters in diameter. Tradeoffs between size of the experimental unit and replication is an important design consideration. Carpenter et al. (1998) conclude that it may be more informative to increase the number of treatments rather than to replicate separate treatments. Tile key question of fundamental concern in the design of these experiments is: How do ecosystems respond non-randomly to physical, chemical or biological manipulation? In a series of very influential papers, Schindler (1988, 1990) and Levine and Schindler (1992) compared the results from long-term ecological experiments performed in whole lakes with results from lake subsamples contained in smaller enclosures, ranging from liter-scale bottles to cubicmeter-scale mesocosms. Unfortunately, the results from these two sets of protocols oftentimes do not track each other. Schindler (1998) has recently reviewed the probable reasons for this ecological mismatch, including physical and biological shortcomings of the subsampled scales. The principal conclusion from these observations is that the whole ecosystem results must be correct, so this is the standard for subsample comparison. While the major limitation in these whole lake experiments is the identification of an acceptable 'twin' for replication treatment, the ocean, 28
because of its large scale, provides an opportunity for replication of a defined ecological unit. Nevertheless, 'whole ecosystem' studies, including manipulation experiments, are rare in the marine environment. The Controlled Ecosystem Pollution Experiment (CEPEX) employed 10 m x 3 m plastic enclosures as experimental marine 'plots' to study the response of marine communities, from bacteria to fish, to the addition of nutrients, trace metals and petroleum hydrocarbons (Menzel and Case, 1977). Recently, oceanographers have used unenclosed, ecosystem-level fertilization to test the hypothesis that primary productivity in selected high-nutrient, low-chlorophyll regions of the world ocean is Fe-limited (Martin et al., 1994; Coale et al., 1996). These perturbations typically lasted for a period of several weeks, at most, so long-lived changes including succession and negative/positive feedbacks were probably not well documented. Nevertheless this 'whole ecosystem' experimental approach is probably the most direct and most relevant for understanding ecosystem dynamics and for improving our prediction of the response of the ocean to natural and anthropogenic change (Carpenter et al., 1995).
e~,~,~,~,e T H E H O T P R O G R A M P R O T O C O L S : A C A S E STUDY Background
Since October 1988, scientists with the Hawaii Ocean Time-series program (HOT) have been evaluating the temporal variability of physical and biogeochemical processes in the subtropical North Pacific Ocean. Our primary sampling site is the deep-water Station ALOHA (A Long-term Oligotrophic Habitat Assessment; 22o45 ' N, 155 ° W). This open ocean site lies approximately 100 km north of the island of Oahu, upwind of the Hawaiian island chain and over flat topography, with a water depth of 4750 m (Karl and Lukas, 1996). The waters around Station ALOHA are considered to be representative of the North Pacific subtropical gyre (NPSG), which is the largest circulation feature on Earth and our planet's largest contiguous biome (Karl, 1999). The water column of the NPSG hosts a vast (in area and depth), low-biomass, microbially-dominated ecosystem that exhibits changes on a variety of timescales. Meaningful sampling of this habitat requires a robust, interdisciplinary field program and modern oceanographic sampling methods.
Site selection
As discussed above, effective sampling of marine ecosystems requires careful selection of sampling locations. While spatial studies of microbiological oceanographic processes may rely on sampling transects or grids for adequate areal coverage, long-term time-series programs typically visit a specific site at regular intervals over an extended period of time. In this model the sampling is Eulerian rather than Lagrangian and this is 29
clearly a critical and debatable sampling strategy. No single site is perfectly representative of the entire oceanic province under study, but the time-series researcher must select a site that is as representative as possible, while keeping in mind the considerable logistic constraints associated with field oceanography. In selecting Station ALOHA for our research, we maintained the absolute criteria that the station must be in deep water (>4000 m), upwind (NNE) of the Hawaiian island chain, and sufficiently far from land to be free from coastal ocean dynamics and terrestrial biogeochemical influences. Within these absolute requirements, we chose a site that would be close enough to the port of Honolulu to make regular (approximately monthly) visitation logistically and financially feasible. At the beginning of each cruise, we also visit the near-shore Station Kahe (21020.6' N, 158016.4' W) to collect comparative data and to test equipment before proceeding to the open-ocean site.
High-resolution depth profiles HOT cruises are conducted approximately monthly and are typically 4 days in duration, with 60-72 h spent at Station ALOHA. This frequency of station occupation allows for some resolution of seasonal and interannual variability in ecosystem parameters. Data collected from moored inst~Tuments (see below) helps to fill in the gaps. During each occupation of Station ALOHA, repeated CTD hydrocasts to 1000 m are performed at approximately 3 h intervals for at least 36 h, in order to assess the physical characteristics of the water column. These data are used to produce a cruise-average density profile, which becomes useful for filtering out high-frequency fluctuations in the depth-density structure of the water column caused by tides and internal waves. The relative depths of biogeochemical features such as particle maxima and nutrient gradients can then be directly compared between cruises. At least one deep hydrocast per cruise is also carried out, to within a few meters of the bottom.
Discrete depth measurements During each monthly station occupation, whole water samples are collected at discrete depths using a 24-place rosette with 12 1PVC bottles (see above). These bottles utilize teflon-coated stainless steel springs and silicone o-rings for closure to minimize sample contamination with leached chemicals. From these primary samples, subsamples are drawn for a suite of chemical and microbiological measurements (Table 2.1). The detailed protocols for processing, storage, and analysis of these subsamples are beyond the scope of this chapter, but are available online at the HOT program website (http://hahana.soest.hawaii.edu/hot/hot_jgofs.html). For most discrete water column measurements, we collect samples at fixed depths from surface to bottom, but the sample spacing is wider in the deep ocean and compacted in the euphotic and shallow aphotic zones (0-250 m). This uneven spacing reflects the decreasing distribution of microbial biomass and biogeochemical variability with depth. We collect 30
Table 2, I Core parameters measured in the Hawaii Ocean Time-series program Parameter
Sensor or analytical procedure Depth or depth range (m)
Continuous profiles Depth (pressure) Temperature (in situ) Salinity (conductivity) Dissolved oxygen Fluorescence PAR and spectral irradiance Natural fluorescence
0-4800 0-4800 0-4800 0-4800 0-1000 0-150 0-150
Pressure transducer on SeaBird CTD Thermistor on SeaBird CTD Conductivity sensor on SeaBird CTD YSI polarographic sensor on SeaBird CTD Sea-Tech fluorometer on SeaBird CTD Biospherical Instruments, PRR-600 Biospherical Instruments, PRR-600
Discrete water bottle samples Salinitv Dissolved oxygen Dissolved inorganic carbon Alkalinity Dissolved nitrate and nitrite Soluble reactive P
0-4800 0-4800 0-4800 0-4800 0-4800 0 4800
Soluble reactive Si Dissolved organic C Dissolved organic N and P Particulate C and N
0-4800 0-1000 0-1000 0-1000
Particulate P
0-1000
Pigments, chlorophyll a
0-200
Primary production Bacteria and cyanobacteria Respiration Bacterial production
0-200 0-200 0-200 0-200
Conductivity Automated Winkler titration Coulometry Automated Gran titration Chemiluminescence and autoanalyzer MAGIC, spectrophotometry and autoanalyzer Autoanalyzer HTCO, IR detection UV digestion, autoanalyzer High-temperature combustion, gas chromatography High-temperature ashing, spectrophotometry High-pressure liquid chromatography and fluorometry 'Clean' HC ill situ incubations Flow cytometry Incubation, Winkler O~ determination Incubation, 'H-leucine uptake
Free-drifting sediment traps Particulate C, N and P Particulate calcium carbonate Particulate biogenic silica
150 150 150
As above Acidification, IR analysis of CO. Alkaline digestion, spectrophotometry
Bottom-moored sequencing sediment traps Particulate C, N and P Particulate calcium carbonate Particulate biogenic silica
1500,2800,4000 1500,2800,4000 1500,2800,4000
As above As above As above
Net tows Meso- and macrozooplankton
0 150
C, N, mass, identification, gut pigments
HALE A L O H A mooring Meteorological measurements
Surface
Dissolved gases Nutrients Optics Temperature
50 120,180 25 0-200
Thermistor, anemeter, pyranometer, rain gauge GTD (Gas Tension Device) Osmoanalyzer Spectral radiometer Thermistor
31
additional samples at the depths of specific hydrographic features, such as the shallow salinity maximum and the deep oxygen minimum. Our sampling depths below the euphotic zone typically coincide with discrete, physically and chemically defined water masses. Also, we collect 'samples of opportunity' from depth strata in which our real-time instruments reveal interesting or unusual phenomena.
Flux and rate measurements
The rate of primary photosynthetic production is a critical parameter for the determination of energy and carbon flows through oceanic ecosystems. We measure the depth profile of planktonic carbon assimilation on each HOT cruise, using 14C tracer methodology. Because of the potential depression of production rates in incubated water samples due to the toxicity of contaminating trace metals, ultra-clean techniques are utilized for these measurements (Fitzwater et al., 1982). A dedicated winch with a kevlar line is used to collect water samples at eight depths from 5 to 175 m, using special Go-Flo ~sampling bottles. These bottles are teflon-coated, and are deployed in a closed position so as to prevent contamination from compounds concentrated within the surface microlayer. Subsamples are drawn into acid-cleaned polycarbonate bottles and spiked with )*Clabeled bicarbonate, then attached to a free-floating array at the depths from which they were originally sampled. The Go-Flo ~ cast is conducted at night to avoid light shock to the organisms, and the array is deployed from the ship at dawn and recovered again at dusk. The incorporation of '4C label into particulate matter during this in situ incubation serves as an estimate of the net photosynthetic fixation of carbon during the daylight hours. The gravitational flux of particles from the euphotic zone is another key rate measurement carried out regularly at Station ALOHA. This sinking material is not only the primary export term for organic carbon and other bioelements, it is also the primary source of energy for the metabolism of mid-water and deep-sea communities. In order to quantitatively estimate this downward flux, we collect particles using a free-drifting sediment trap array, following the Multitrap design (Knauer et al., 1979). The particle interceptor traps (PITs) are cylindrical polycarbonate tubes containing a high-density formalin-amended salt solution, which serves both to prevent wash-out and to preserve the collected particles. Twelve PITs at each depth are attached to the array line with a PVC cross-bar, and the array is left to float freely for 60-72 h. We routinely deploy these sediment traps near the base of the euphotic zone (150 m) to catch the euphotic zone export flux, and at other depths as desired for additional information about particle decomposition dynamics.
P l a n k t o n n e t tows
If we are to acquire a holistic understanding of microbial dynamics in the sea, we must include studies of microbial mortality and the passing of 32
carbon and energy to higher trophic levels, in the HOT program, the community structure of meso- and macrozooplankton is examined through the routine collection of these larger organisms using towed plankton nets. This method of sampling is not only useful for collecting zooplankton grazers, but it is also effective for collecting the rare large (>64 lJm) microalgal and cyanobacterial cells, and in particular the colonies and aggregates thereof.
Light and meteorology Biological processes in the upper ocean are strongly affected by weather, for example, through the influence of wind on water column mixing, and through the influence of light on photosynthesis. Accordingly, we routinely collect data on incident solar irradiation and spectral radiance with depth in the upper water column (0-175 m), as well as a suite of shipboard meteorological measurements (Table 2.1).
Moored instruments Because monthly sampling cannot effectively resolve short-term (minutesweeks) variability of ocean biogeochemistry, we utilize electronic datacollecting instruments attached to a moored buoy system as much as possible. The HALE ALOHA mooring (Hawaii Air-sea Learning Experiment at Station ALOHA) is deployed at a fixed position near the station for several months at a time. The mooring is outfitted with a variety of devices, including CTDs, optical sensors, dissolved gas sensors, nutrient analyzers and meteorological instruments. Although not a substitute for hands-on microbiological sampling and experimentation, the highfrequency data sets obtained from the moored instruments provide an unprecedented window on the environmental variability in the NPSG, and complement the monthly sampling scheme. Details of the instrumentation used on HALE ALOHA can be found on the HOT program website (http://hahana.soest.hawaii.edu/hot/hale-aloha/ha.html).In addition to HALE ALOHA, we have repeatedly deployed an array of time-series sequencing sediment traps. These bottom-moored instruments collect the flux of sinking particles reaching the deep ocean (4000 m) over weekly intervals, and thus provide a detailed export data set complementary to that obtained from the monthly free-floating sediment trap deployments.
Ancillary measurements and experiments The core physical and biogeochemical data collected by HOT program scientists represent more than just an exercise in long-term microbial habitat monitoring - - they represent a framework upon which a conceptual and mechanistic understanding of ocean ecosystem dynamics may be built. To that end, the HOT program has hosted a wide array of collaborating researchers, whose measurements and experiments complement 33
our own efforts. These ancilliary projects have included investigations of phytoplankton population dynamics, microbial production of trace gases, isotopic constraints on the ocean's role in the global carbon cycle, and molecular biological evaluations of nitrogen fixing genes, to name but a few. A list of these collaborations may be found on the HOT program (http://hahana.soest.hawaii.edu/hot/ancillary.html). The website success of each of these endeavors relies, ultimately, upon the proper collection of appropriate samples.
CONCLUSION
AND PROSPECTUS
Seagoing microbiological oceanographers labor under several disadvantages compared to their land-based counterparts. First and foremost is the immense scale of the habitat, relative to the size of the sample that is generally removed for quantitative analysis. In addition, it is no longer reasonable or acceptable for marine microbiologists to focus exclusively on the microbial inhabitants of the sea, especially if in situ ecological process understanding is a primary objective of the investigation so great care must be given to collections and analyses of complementary physical and biogeochemical data sets. Second is the difficulty in conducting controlled, replicated field experiments. Finally, the complexity and ephemeral nature of most marine ecosystems precludes straightforward scaling from the experimental measurements to regional or basin-wide scales with any degree of certainty. Empirical modeling approaches, that have been used extensively in ecological studies, must be supplemented with process-oriented studies of the physical, chemical and biological interactions before a comprehensive understanding can be achieved. High on the relatively long list of research priorities is the design and implementation of a rigorous sampling program. It is dangerous to predict the future progress of any scientific discipline. tn the case of microbiological oceanography, however, there is ample evidence to suggest that the field will experience a rapid increase in technology and instrumentation, including sampling, over the next few decades. Field researchers should anticipate increased automation both in sample collection and in the remote, autonomous detection of microorganisms (including specific microbes) and their in situ metabolic activities. Futhermore, advanced statistical treatments of ecological data and the development of comprehensive models should provide better understanding of the complex interactions between microorganisms and their environment. It is very likely going to be an exciting few decades.
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3 Enumeration of Viruses Rachel T N o b l e Southern California CoastaIWater Research Project (SCCWRP), 7171 Fenwick Lane,Westminster, CA, 92683 USA
CONTENTS Introduction Method Troubleshooting Applications Conclusions
~,~,~,~,~,~, I N T R O D U C T I O N Viruses are now known to be the most numerically abundant component of marine plankton (Bergh et al., 1989; Bratbak et al., 1990; Fuhrman and Suttle, 1993; Hennes and Suttle, 1995; Noble and Fuhrman, 1998; Fuhrman, 1999). In the late 1980s, some of the first reports were published documenting the high abundances of marine viruses at 10 '~' particles per liter of seawater, exceeding the typical abundance of bacteria (Proctor et al., 1988; Bergh et al., 1989). Since then, studies have demonstrated high numbers of viruses in all types of marine environments, from eutrophic coastal waters to deep blue open-ocean waters, from the sea surface to the depths of the sea, and from the polar to the tropical regions (Bratbak et aI., 1990; Cochlan et al., 1993; Guixa-Boixareu et al., 1996; Steward et al., 1996). Multiple groups of researchers have identified important roles of viruses in the mortality of heterotrophic bacterioplankton, cyanobacteria, and phytoplankton. They also play a role in biogeochemical cycling and control of species diversity (Fuhrman, 1999; Wilhelm and Suttle, 1999). Specifically, it has been shown by a number of researchers that viruses are capable of causing a significant portion of the heterotrophic bacterial mortality in certain marine environments (Fuhrman and Noble, 1995; Guixa-Boixareu et al., 1996; Steward et al., 1996; Weinbauer and H6fle, 1998). Current research in the fields of marine microbiology and marine microbial ecology requires the ability to rapidly enumerate viruses and bacteria. In the past, counting microbes in seawater samples by transmission electron microscopy (TEM) was the standard method (Bergh et al., 1989; Borsheim et al., 1990). This method is tedious, expensive, involves METt tODS IN MICROBIOLOGY, VOLUME 30 ISBN 0 12 521530 4
Copyright © 2001 Academic Press Lid All rights of reproduction in any form reserved
time-consuming preparatory steps, lacks precision, and requires expensive ultracentrifugation and electron microscopy equipment not available to many researchers. In recent years, other stains such as DAPI (4'6diamidino-2-phenylindole) and Yo-Pro I (Molecular Probes, Inc.) have been used for enumeration of virus particles by epifluorescence microscopy (Suttle et al., 1990; Hara et al., 1991; Proctor and Fuhrman, 1992; Hennes and Suttle, 1995; Weinbauer and Suttle, 1997; Xenopolous and Bird, 1997). However, DAPI is not sufficiently bright to be used with direct visual observation on many microscopes. Therefore, it has been necessary for some researchers to use photomicrography or image intensification in order to count and size virus particles (Hara et al., 1991; Fuhrman et al., 1993). Newer microscopes may allow direct visual counts with this stain, but many labs do not possess high-powered microscopes and have turned to the use of brighter stains (Weinbauer and Suttle, 1997). Yo-Pro I has recently been used for seawater studies (Hermes and Suttle, 1995; Weinbauer and Suttle, 1997). The stain intensity is bright, but the stain is not compatible with aldehydes (such as formaldehyde and glutaraldehyde), it requires extra dilution and rinsing steps to remove salts, and the staining time is 2 days. Improvements have been made on the Yo-Pro I method originally reported by Weinbauer and Suttle (1997), which involved microwaving of the samples to permit penetration of the stain (Xenopolous and Bird, 1997). A newer stain, SYBR Green I, has been developed. The aim of this chapter is to provide the details necessary for enumeration of viruses and bacteria in seawater using the nucleic acid stain SYBR Green I (Molecular Probes, Inc.). This stain was originally used for research using flow cytometry by Marie et al. (1997). SYBR Green I is a viable tool which yields virus counts comparable to TEM in a broad variety of samples, and seems to be more easily applied to the analysis of seawater samples than some of the previously mentioned stains. SYBR Green I has the advantages of being usable in conjunction with seawater and commonly used fixatives and a short staining period. SYBR Green I stained viruses and bacteria are intensely stained and easy to distinguish from other particles with both older and newer generation epifluorescence microscopes. In addition to the methodological advantages that SYBR Green I offers, it is inexpensive and its manufacturer claims it to be less carcinogenic than other typical nucleic acid stains. It has recently been noted that another nucleic acid stain, SYBR Gold, also made by Molecular Probes, Inc., can be used interchangeably with SYBR Green I. This stain appears to require a slightly shorter staining time (12 min), and is less expensive. Although the author has not performed a quantitative comparison between SYBR Green I and SYBR Gold, it appears that this latter stain can be used interchangeably with the protocol listed here, with the minor change in the staining time (Chen et al., in press).
44
ENUMERATION GREEN I
OFVIRUSES
USING SYBR
Principle The SYBR Green I m e t h o d is used for easy and rapid enumeration of both marine viruses and bacteria in seawater samples. Seawater samples are collected and fixed with formalin, which cross-links the proteins found in cell m e m b r a n e s and viral coat proteins. Marine bacteria and viruses are very a b u n d a n t in seawater, sufficient that w h e n seawater samples are filtered with an ultra-fine pore size filter (Anodisc, 0.02 ~m), they can be counted by epifluorescence microscopy. The material on the filter can be stained with a variety of stains, but for quick and inexpensive viral and bacterial counts, we have found the nucleic acid stain SYBR Green I to be the most advantageous. By diffusion, the stain permeates the filter and stains any particles containing DNA or RNA. Convenient quantification can be achieved by m o u n t i n g the filter on a glass slide, with the use of an anti-fade m o u n t i n g solution u n d e r the cover slip, and counting tile fluorescent particles by epifluorescence microscopy. The average n u m b e r of viruses and bacteria counted per r a n d o m l y selected field is multiplied by a conversion factor which represents the total n u m b e r of microscope fields that fit into the total available filter area. This conversion factor is then used to calculate a total n u m b e r of viruses or bacteria per filtered volume, often expressed as virus particles or cells ml ', respectively.
Equipment and reagents • • • • • • • • • • •
Formalin (37 % formaldehyde solution, Sigma Chemical, Inc.) 50 ml polypropylene conical tubes for sampling (Fisher Scientific, Inc.) Whatman Anodisc 0.02 ~m Membrane Filters (Fisher Scientific, Inc.) Millipore 0.8 ~m filters (Fisher Scientific, Inc. or Millipore, Inc.) Filtration manifolds and towers (Fisher Scientific, Inc.) SYBR Green I stain (Molecular Probes, Inc.) Sterile, 0.02 ~m filtered, deionized water Plastic Petri dishes (VWR Scientific Inc.) Eppendorf pipets and tips (VWR Scientific, Inc.) Slides and cover slips (VWR Scientific, Inc.) Mixture of 50% Phosphate Buffered Saline (0.05 M Na2HPO4, 0.85% NaCI, pH 7.5, Sigma Chemical Co.) and 50% glycerol (Sigma Chemical Co., should be made in advance and stored in the refrigerator) • p-Phenylenediamine (dihydrochloric acid, 10% w/v, Sigma Chemical Co., should be made in advance and stored frozen, in the dark)
Assay Sample collection 1. Collect seawater samples using Niskin bottles or triple acid-rinsed bottles (5 ~ hydrochloric acid), or by bucket and transferred into
45
acid-rinsed and sample-rinsed 50 ml polypropylene tubes (Fisher Scientific, Inc.) . A d d 0.02 ~m filtered formalin to the sample(s) to a final concentration of 1%. If the samples are not going to be filtered immediately, they should be stored in the dark at 4°C. Fixed samples can be stored chilled for up to a week, but optimally should be processed as soon as possible. Slide preparation 1. Whenever possible, preparation should be done under subdued light (dimmed room light is suitable). When ready to begin, perform a dilution of SYBR Green I stock from Molecular Probes, Inc. to 1:10 of the supplied concentration with 0.02 ~m filtered, sterile, deionized water. For example, dilute 5 ~1 of the SYBR Green I stock solution with 45 ~1 H20. Put unused stock immediately back at -20°C. Immediately prior to sample filtration prepare the anti-fade mounting solution. To do this, mix 990 ~tl of the 50% PBS/50% glycerol mixture with 10 ~tl of 10% p-phenylenediamine. Store this solution on ice, in the dark, while working. When removing the 10 ~tl of p-phenylenediamine, thaw, vortex, remove the 10 ~1 and then refreeze immediately. Also, for each filter to be stained, place a 97.5 ~1 drop of 0.02 ~m filtered, sterile, deionized water inside a clean plastic Petri dish. To each drop of water, add 2.5 ~I of the 10% SYBR Green I working solution (final dilution 2.5 x 10 ~). Keep the Petri dishes with the drops of stain in a cool, dark place during the course of the filtration. Filter a I to 10 ml formalin-fixed seawater sample through a 25 mm, 0.02 ~m pore-size, Anodisc membrane filter (Fisher Scientific, Inc.), backed by a 0.8 ~m cellulose mixed ester membrane (type AA, Millipore, Inc.) at 15-20 kPa vacuum. The volume of seawater to be filtered depends upon the type of seawater used, eutrophic estuarine samples will require filtration of only about 1 ml, whereas open-ocean seawater samples m a y require filtration of up to 10 ml in order to provide for statistically meaningful bacterial and viral counts. Filter the Anodisc to dryness and remove it with forceps with the vacuum still on. Lay the filter, sample side up, on a drop of the staining solution in the Petri dish for 15 rain in the dark. After the staining period, pick the filter up with forceps and carefully wick away any remaining moisture by touching the back side of the membrane to a Kimwipe (any droplets on the top plastic rim of the filter should also be blotted). 6. Place the filter on a glass slide. Onto a 25 mm square cover slip, place a 30 ~1 drop of the anti-fade mounting solution (50% PBS / 50% glycerol with 0.1% p-phenylenediamine). Invert the cover slip, drop-side down, onto the filter. Press d o w n on the cover slip with a 2.
.
.
.
46
Kimwipe to be sure that all bubbles are displaced. If the SYBR Green I stained filters are to be counted immediately, place a drop of immersion oil on top of the cover slip. 7. For reading the slides, r a n d o m l y select 10-20 fields to count a total of >200 viruses and >200 bacteria per filter u n d e r blue excitation on an epifluorescence microscope equipped with a 100 W Hg lamp. Virus particles will appear as distinctly shaped 'pinpricks' and fluoresce bright green, and bacterial cells will be much brighter and should easily be distinguished from viruses because of their relative size.
Troubleshooting SYBR Green I slides should be counted immediately, but can be stored frozen for 2-3 weeks. W h e n preparing the anti-fade m o u n t i n g solution, r e m e m b e r that p - p h e n y l e n e d i a m i n e is quickly oxidized at room temperature. Tubes of p - p h e n y l e n e d i a m i n e can be t h a w e d / f r o z e n only about three times before they need to be discarded. If a b r o w n color is noted in the p - p h e n y l e n e d i a m i n e solution, discard it, and immediately make a fresh solution. If, w h e n the slides are being counted, the viruses appear to be in more than one focal plane, or appear to be floating, remake the slide. Also, if the background fluorescence makes it difficult to count the slide, i.e. the slide appears washed out, remake the slide, staining the filter for the prescribed a m o u n t of time.
Applications Seawater samples stained with SYBR Green I and observed u n d e r an epifluorescence microscope demonstrate bacteria that are intensely stained, and virus particles that are brightly stained and countable (Plate 1). Detritus has not been significantly stained by SYBR Green I in past observations. Previous analysis of coastal samples in Noble and Fuhrman (1998), d e m o n s t r a t e d bacterial counts by SYBR Green I that were essentially identical to acridine orange counts, with an r -~of 0.99. Virus counts by both SYBR Green I and TEM are highly correlated (r: = 0.93, p < 0.001, Figure 3.1). There is a tendency for tile SYBR Green I counts to be higher, as indicated by the slope of the linear regression being 1.10 (Figure 3.1). In Noble and F u h r m a n (1998), virus counts by SYBR Green I and TEM s h o w e d very similar patterns in seawater samples from all depths, with SYBR Green I counts about 25% higher than TEM counts. Freshwater samples stained with SYBR Green I demonstrated viruses and bacteria that appear to be even more intensely stained than those from seawater (Fuhrman and Noble, 1998). In recent studies by Hermes and Suttle (1995) and Weinbauer and Suttle (1997), Yo-Pro 1 based virus counts were found to average about 2.3 and 47
3.0
C
E
2.0
"J
x
a3 1.0
•
(3 n>-
•
0.00.0
•
1.0
i
2.0
3.0
TEM (x 10 7 virus ml "1)
Figure 3.1. Comparison of virus counts using SYBR Green I and transmission electron microscopy (TEM) for a diverse set of marine samples. Error bars indicate the standard deviation of duplicate samples; where they are not seen, the standard deviation was less than the size of the symbol. Line indicates linear regression (r2 = 0.92, a subset of the data used for this graph appears as part of Figure 3 in Noble and Fuhrman, 1998).
1.5 times higher than counts by TEM, respectively, over a wide range. It has been demonstrated that at lower viral densities, TEM counts were generally similar to those by SYBR Green I, and at higher viral densities, TEM counts were clearly lower than those for SYBR Green I (Noble and Fuhrman, 1998). This trend is also consistent with work published by both H e n n e s and Suttle (1995), and Weinbauer and Suttle (1997) w h e n comparing Yo-Pro I and TEM. Both of these studies have also m a d e comparisons with an alternative stain for epifluorescence microscopy, DAPI. However, DAPI is relatively dim, and requires high-quality optics for quantitative visualization of viruses. SYBR Green I can be used to stain virus and bacterial particles in m a n y different types of samples, marine, freshwater, and sediment included. It is apparently not inhibited by the use of fixatives, the staining period is short, and SYBR Green ! is reported to stain both RNA and DNA viruses. In certain environments, it m a y be necessary to increase the recomm e n d e d concentration of SYBR Green I (2.5 x 10 ~ dilution) to yield brighter and more stably fluorescent viruses. Fading of the samples is best retarded with the use of the prescribed anti-fade m o u n t i n g solution. SYBR Gold (Molecular Probes, Inc.) is another sensitive fluorescent stain for detecting double- or single-stranded DNA or RNA. According to the product information from Molecular Probes, Inc., SYBR Gold has been s h o w n to be more sensitive than SYBR Green [ and II for staining nucleic acids in electrophoresis. A parallel comparison between SYBR
48
Gold and SYBR Green I was recently carried out by staining the same seawater sample (Chen et al., in press). It was found that the epifluorescence signal of SYBR Gold stained viruses lasted longer than that of SYBR Green I stained viruses w h e n the final concentration (2.5x) was used for both SYBR stains. Without using any anti-fade m o u n t i n g solution, the fluorescence of SYBR Gold stained viruses was stable for more than 2 min, while the SYBR Green I signal faded within 30 s. When a higher concentration (25x) of SYBR Gold or Green I was used, some bacterial cells in natural samples appeared to be overstained and their fluorescent halos could overcast the fluorescent signal of stained viruses. SYBR Gold is a less expensive nucleic acid stain than SYBR Green I, and can be a good alternative fluorochrome for fast staining and accurate counting viral particles in various types of aquatic samples. Based on these results, the author r e c o m m e n d s the use of the presented protocol with either SYBR Green I or SYBR Gold in conjunction with 0.02 ~m pore size Anodisc filters for determining the viral and bacterial abundance in seawater. Nucleic acid stains such as SYBR Green I and SYBR Gold m a y be used increasingly in the future for more efficient approaches, such as automated counting of viruses. Chen et al. (in press) have recently also used digital image analysis and flow cytometric analysis for rapid counting of SYBR Gold stained viral particles. Flow cytometric analysis s h o w e d that fluorescence per virus stained with SYBR Gold was about 2 times higher than that stained with SYBR Green I (Chen et al., in press). They also found that digital image analysis could detect some weakly stained viruses in natural samples that could not easily be detected by the h u m a n eye. A further advantage of the use of fluorescent stains is that digital images of both SYBR Gold and SYBR Green I stained samples can be quickly captured and saved in the c o m p u t e r for later processing. In the past, it has been quite difficult to incorporate viruses and virusmediated processes into research on aquatic food webs. This m e t h o d allows reasonably e q u i p p e d microbiology laboratories to perform rapid counts of virus particles in natural samples. Virus counts with SYBR Green I or SYBR Gold can be p e r f o r m e d easily in the lab or on board ship and m a y help elucidate the roles of viruses in aquatic systems.
References Bergh, O., Borsheim, K. ¥., Bratbak, G. and Heldal, M. (1989). High abundance of viruses found in aquatic environments. Nature 340, 467-468. Borsheim, K. ¥., Bratbak, G. and Heldal, M. (1990). Enumeration and biomass esfimarion of planktonic bacteria and viruses by transmission electron microscopy. Appl. Environ. Microbiol. 56, 352-356. Bratbak, G., Hetdal, M., Norland, S. and Things{ad, T. E (1990). Viruses as partners in spring bloom microbial trophodynamics. Appl. Environ. Microbiol. 56, 1400-1405. Chen, E, Lu, J. R., Binder, B. J., Liu, Y. C. and Hodson, R. E. (2000) Enumeration of marine viruses stained with SYBR Gold: Application of digital image analysis and flow cytometry. Appl. Environ. Microbiol. M press. 49
Cochlan, W. P., Wikner, J., Steward, G. E, Smith, D. C. and Azam, E (1993). Spatial distribution of viruses, bacteria and chlorophyll a in neritic, oceanic and estuarine environments. Mar. Ecol. Prog. Ser. 92, 77-87. Fuhrman, J. A. (1999). Marine viruses and their biogeochemical and ecological effects. Nature 399, 541-548. Fuhrman, J. A. and Noble, R. T. (1995). Viruses and protists cause similar bacterial mortality in coastal seawater. Limnol. Oceanog. 40, 1236-1242. Fuhrman, J. A. and Suttle, C. A. (1993). Viruses in marine planktonic systems. Oceanography 6, 51-63. Fuhrman, J. A., Wilcox, R. M., Noble, R. T. and Law, N. C. (1993). Viruses in marine food webs. ln: Trends in Microbial Ecology (C. Pedros-Alio and R. Guerrero, Eds) Spanish Society for Microbiology: Barcelona, Spain), pp. 295-298. Guixa-Boixareu, N., Calderon-Paz, J. I., Heldal, M., Bratbak, G. and Pedros-Alio, C. (1996). Viral lysis and bacterivory as prokaryotic loss factors along a salinity gradient. Aquatic Microbial Ecol. 11, 215 227. Hara, S., Terauchi, K. and Koike, I. (1991). Abundance of viruses in marine waters: Assessment by epifluorescence and transmission electron microscopy. Appl. Environ. MicrobioI. 57, 2731-2734. Hennes, K. P. and Suttle, C. A. (1995). Direct counts of viruses in natural waters and laboratory cultures by epifluorescence microscopy. Limnol. Oceanog. 40, 1050-1055. Hennes, K. P., Suttle, C. A. and Chan, A. M. (1995). Fluorescently labeled virus probes show that natural virus populations can control the structure of marine microbial communities. Appl. Environ. Microbiol. 61, 3623-3627. Marie, D., Partensky, E, Jacquet, S. and Vaulot, D. (1997). Enumeration and cellcycle analysis of natural-populations of marine picoplankton by flow-cytometry using the nucleic-acid stain SYBR green I. Appl. Environ. Microbiol. 63, 186-193. Noble, R. T. and Fuhrman, J. A. (1998). Use of SYBR Green i for rapid epifluorescence counts of marine viruses and bacteria. Aquatic Microbial Ecol. 14, 113-118. Proctor, L. M. and Fuhrman, J. A. (1992). Mortality of marine bacteria in response to enrichments of the virus size fraction from seawater. Mar. Ecol. Prog. Ser. 87, 283-293. Proctor, L. M., Fuhrman, J. A. and Ledbetter, M. C. (1988). Marine bacteriophages and bacterial mortality. EOS Trans. Am. Geophys. Union 69, 1111-1112. Steward, G. E, Smith, D. C. and Azam, E (1996). Abundance and production of bacteria and viruses in the Bering and Chukchi Sea. Mar. Ecol. Prog. Ser. 131, 287 300. Suttle, C. A., Chan, A. M. and Cottrell, M. T. (1990). Infection of phytoplankton by viruses and reduction of primary productivity. Nature 387, 467-469. Weinbauer, M. G. and H6fle, M. G. (1998). Size-specific mortality of lake bacterioplankton by natural virus communities. Aquatic Microbial Ecol. 15, 103-113. Weinbauer, M. G. and Suttle, C. A. (1997). Comparison of epifluorescence and transmission electron microscopy for counting viruses in natural marine waters. Aquatic Microbial Ecol. 13, 225-232. Wilhelm, S. W. and Suttle, C. A. (1999). Viruses and nutrient cycles in the sea. Bioscieilces 49, 781-788. Xenopolous, M. A. and Bird, D. E (1997). Virus a la Sance Yo-Pro: Microwave enhanced staining for counting viruses by epifluorescence microscopy. Limnol. Oceano% 42, 1648-1650.
50
List of suppliers Fisher Scientific, Inc. 585 Alpha Drive Pittsburgh, PA 15238, USA Teh 1 800 766 7000 Fax: 1 800 926 1166 http://www.fishersci.conl
Millipore filters, filtration supplies, Anodiscs
VWR Scientific Products, Inc. V W R International 3000 Hadley Road So. Plainfield, NJ 07080, USA Teh 1 800 932 5000 07" 1 908 757 4045 Fax: 1 908 757 0313 http://www.vwrsp.com
Anodisc filter membranes, slides, cover slips
s._ °~
0 0 4,a
°~
Molecular Probes, Inc. 4809 Pitchfi~rd Avettue Eugene, OR 97405-0469, LISA T~q: 1 541 465 8300 Fax: 1 541 344 6504 h t t p :/ /www.probvs.c om
SYBR Green I, SYBR Gold Sigma Chemical Co. P.O. Box 14508 St Louis, M O 63178, USA Teh 1 800 325 3010 or 1 314 771 5750 Fax: 1 800 325 5052 or 1 314 771 5757
p-Phenylenediamine (dihydrochloride), disodium phosphate for PBS
51
Whatman Lab Sales St Leotlard's Road 20/20 Maidstone, Kent ME16 OLS Telephone: +44(0)1622 674821 Fax: +44(0)1622 682288 E-maih labsales@whatmm~.com Whatman h~ternational Ltd., Cataloxue Sales Departme~#
Direct purchase of Anodisc filters
0~ L
E ILl
4 Quantification of Algal Viruses in Marine Samples StevenWWilhelm and Leo Poorvin Department of Microbiology, The University of Tennessee,Knoxville,TN 3 7996, USA
CONTENTS Introduction and background Concentration of viruses in water samples by ultrafiltration Most probable number (MPN) assays Plaque assays Conclusions
~HI,~I,41,~HI, I N T R O D U C T I O N
AND
BACKGROUND
Phycoviruses (viruses that infect either cyanobacteria or eukaryotic algae) impart significant mortality on their hosts in aquatic environments. Microorganisms (both eukaryotic and prokaryotic) in marine systems are thought to be responsible for as m u c h as 50% of the photosynthetic carbon fixation on the planet (Field et al., 1988). It is therefore apparent that agents of mortality that act directly to reduce primary production in marine environments will alter carbon and energy flux through these systems (Wilhelm and Suttle, 1999; Fuhrman, 1999). This has, in part, led to the increased interest in the ecology of marine viruses that has occurred through the last decade. Studies concerning the distribution and activity of viruses at the c o m m u n i t y level c o m m o n l y rely on direct counts to monitor changes in the natural viral community. While this information is pertinent to m a n y studies, it does not address the issue of the infectivity of these viruses or the range of organisms that m a y be directly influenced by viral activity. Outside molecular techniques (see below), the identification and enumeration of phycoviruses requires the observation of interactions between virus and their hosts. It is therefore pertinent to m a n y studies to be able to quantify the a b u n d a n c e of infective viruses that m a y impart mortality on specific phytoplankton. However, as these measurements require that virus-host interactions be observed, it is necessary from the onset that the host p h y t o p l a n k t o n be cultivable. Therefore, the techniques highlighted in this paper require that the host organisms can be cultured in the lab in order to enumerate potential viruses. METHODS IN MICROBIOLOGY,VOI~UME30 ISBN (t 12 521530-4
Copyright © 2001 Academic Press I.td All rights of reproduction in any form reserved
The identification of viruses in the sea that infect specific cyanobacteria and bacteria is still in its relative infancy compared to studies on viruses infecting marine heterotrophic prokaryotes (Suttle, 1996). While the total abundance of virus-like particles ranges from 10~ to 10~ m l ' seawater (Wilhelm and Suttle, 1999), viruses infecting and lysing phytoplankton only represent a subset of this population. However, viruses infecting the marine Synechococcus spp. commonly occur at concentrations > 10~ml ' in coastal waters, and have estimated at concentrations as high as 2.5 × 10~ ml' (Suttle and Chan, 1993; 1994; Waterbury and Valois, 1993). Concentrations of viruses infecting eukaryotic phytoplankton can be equally as high; Cottrell and Suttle (1995) measured abundances of lytic viruses infecting Micromonas pusilla at > 10~ ml '. Viruses infecting other phytoplankton, including Aureococcus anophag~{frrens (Milligan and Cosper, 1994), Chrysochromulina spp. (Suttle and Chan, 1995), Emiliania huxleyi (Bratbak et al., 1993), Heterosigma akashiwo (Nagasaki and Yamaguchi, 1997; Lawrence et al., 2000) and Phaeocystis pouchetii (Jacobsen et al., 1996) have also been isolated from pelagic marine systems in recent years. In recent years it has also been demonstrated that infectious phycoviruses can also be isolated from marine sediments. In the Western Gulf of Mexico, Rodda et al. (1996) found cyanophages in concentrations ranging from 9.4 x 10* m l ' at the sediment/water interface of a 47 m water column, to 3.0 x 102 ml ~at 30 cm below the sediment surface. As the water over the sediment contained an order of magnitude less virus, this suggests that the vertical transport and subsequent burial of infectious cyanophage or infected cyanobacteria was occurring (as the production of cyanophage in the absence of light is unlikely). As molecular techniques for the enumeration of phycoviruses are currently under development, it remains premature to include them as protocols in this chapter. Using the polymerase chain reaction (PCR) and virus specific primers, Suttle and co-workers have been able to establish a baseline of information on the genetic diversity of one group of algal viruses, the Phycodnaviridae (Chen and Suttle, 1996; Short and Suttle, 1999). Recently they have been able to estimate the diversity of at least a portion of the Phycodnaviridae using degenerate primers for the segments of the DNA polymerase genes of these algal viruses and denaturing gradient gel electrophoresis. Similarly, Fuller et al. (1998) have described the genetic diversity of cyanophage isolates infecting Synechococcus spp. using PCR techniques targeted at the DNA region encoding a capsid assembly protein. However, as with studies involving viruses infecting eukaryotic phytoplankton, these results remain qualitative. The advent of new techniques (e.g. quantitative PCR, in situ PCR) will hopefully soon provide qualitative values for the distributions of these algal viruses. This review describes the current methods available for the enumeration of specific viruses infecting phytoplankton in aquatic environments. It represents a compilation of methods that have been employed for many years in classic virology and those that have been adapted for use by 'viral ecologists' working in natural systems. Two approaches, the plaque assay $4
and MPN assay, are described here which allow researchers to both e n u m e r a t e and isolate viruses that lytically infect marine photoautotrophs.
C O N C E N T R A T I O N OF VIRUSES IN W A T E R SAMPLES BY U L T R A F I L T R A T I O N Principle In m a n y situations the a b u n d a n c e of lyric phycoviruses in a natural water sample is too low to accurately quantify. In these cases, the use of ultrafiltration techniques m a y be required to increase the concentration of viruses in the sample. Ultrafiltration involves the removal of bacterial and algal c o m p o n e n t s (> 0.2 pro) of the microbial c o m m u n i t y followed by the concentration of the 'viral size fraction' (typically 30 kDa to 0.2 pm). Small scale (0.3-20 ml) ultrafiltrations can be carried out with a commercially available centrifugation systems ('spin-columns') such as Centriprep or Centriplus units (Millipore). While these often will w o r k well with laboratory virus-host systems, these sample sizes are often too small to properly examine environmental samples. In these cases, techniques such as tangential flow filtration or vortex flow filtration can be used to handle larger volumes (1-200 1). With these techniques, concentration of the viral particles is achieved by successive circulation of the sample across a 30 kDa m e m b r a n e surface. This allows the water to be removed from the sample (ultrafiltrate) while the viruses are concentrated into the retained volume. The resulting viral concentrate can then be used as the 'sample' to screen (as described below). In this protocol, we describe the use of the Amicon M12 ultrafiltration system, as this is the system currently in use in our laboratory (adapted from Chen et al., 1996). Similar systems are p r o v i d e d by other suppliers, and it is suggested that the reader consider these other alternatives prior to making any investment in a system.
Equipment and reagents • Submersible pump with pressure gauge. • Two containers for water samples (20-200 I each, depending on volume to be concentrated). • 142 mm diameter glass fiber filters (MSF GC50; nominal pore size, 1.2 IJm) with holder(s) and appropriate tubing (non-toxic). • 142 mm diameter, 0.2 lure nominal pore-size filters (polycarbonate or low protein binding) with holder(s) and appropriate tubing. • Amicon ProFlux M-12 ultrafiltration system with non-toxic tubing designed for use in peristaltic pumps (such as Masterflex from PharMed). • Millipore S IOY30 spiral wound membrane cartridges (30 kDa molecular weight cutoff). • Header kits for S I0 cartridges.
55
Application Collect the water sample (20-200 1) into one of the holding containers. Using the submersible p u m p , prefilter (at < 17kPa) the sample first through the 142 m m diameter glass fiber filters (MSF GC50; nominal poresize, 1.2 ]am). Two or more of these m a y be set up in parallel for larger sample volumes. Follow this by filtering the sample through a 0.2 l~m filter into the second container. These filters will remove large particulates, algae, bacteria, etc. but will allow most viruses to pass. T h r o u g h o u t these steps, subsamples of water should be collected so that the recovery efficiency of this process can be calculated (see below). After filtration, use an Amicon ProFlux M-12 ultrafiltration system to concentrate the filtrate containing the viruses. Set the M-12 u p for concentration mode, with a Millipore $10Y30 ($10) spiral w o u n d m e m b r a n e cartridge (30 kDa cutoff). This cartridge, with a total m e m b r a n e area of 0.93m 2, will allow water to pass through but retain virus particles. Connect tubing from the container with the filtrate into the p u m p inlet of the M-12. Standard operating procedures involve running the p u m p at 40 to 50% of the m a x i m u m speed, with 50 to 60 kPa of backpressure. From the p u m p , run the tubing to the inlet header of the S10 cartridge and from the outlet header back to the container holding the sample to return the retenate. Connect tubing from the permeate connector to remove the ultrafiltrate, which can be discarded. As the system runs, the permeate (without virus particles) is removed, thereby concentrating the viruses in the remaining retenate. With this setup, a v o l u m e of 200 1 of seawater can be concentrated to 500 ml in about 1 hour. Take care not to attempt to reduce the v o l u m e of the retentate below the s u m m e d v o l u m e of the cartridge and the tubing. Measure the final volume of viral concentrate so that the concentration factor (CF) can be estimated as follows:
CF = volume qf sample / volume of retentate If required, the retenate can be further concentrated to a smaller v o l u m e (100 to 200 ml) using a smaller system (e.g. an $1Y30 cartridge, Chen and Suttle, 1996). Alternatively, other low-cost methods are available to concentrate samples. Centrifuge-based concentrators (e.g. Centriprep, Centriplus) can be used to concentrate viruses in small volumes of sample. The recovery efficiency (a.k.a. concentration efficiency) of this process must be determined w h e n using viral concentrates to estimate the abundance of infectious phycoviruses. This can be most easily determined by direct counts of the total viral abundance pre- and post-concentration (see chapter by Noble for direct counting techniques). The recovery efficiency (RF, as a %) is determined as follows:
RF = 100 (A,,,,/A ,,,,) / CF where A , , is the abundance of virus particles in the viral concentrate, A ...... is the abundance of viruses in the original sample, and CF is the concen-
56
tration factor (determined above). Typical recovery efficiencies vary, but are generally greater than 50% and commonly approach 100% (Suttle et al., 1991; Wommack et al., 1995).
Troubleshooting A series of problems can occur when making and using viral concentrates. Most problems are associated with the concentration of the viruses. The problems include leakage from old tubing, loss of viruses during prefiltration, and incorrect estimates of the viral concentration factor. Establishing familiarity with the concentration equipment and procedure(s) is a sure cure for many of these problems. The other considerations to be made with ultrafiltration are problems associated with filter integrity and cleaning. Breakthrough of viruses in damaged filters can seriously hinder concentration efficiency. All manufacturers, however, provide instruction on testing the integrity of their filters. As filters are often costly, cleaning procedures are important and designed to maximize the life of the filter cartridge. In the case of the system described, we recirculate 0.1 M NaOH to remove residual organics from the filter after each concentrate is made. The NaOH is subsequently removed using ddH:O, dilute H~PO~ and then ddH~O. Again, all manufacturers provide information on the chemical compatibility of their filters, and this should be checked in each case. This is especially important as some sterilizing agents (hydrogen peroxide, bleach, strong NaOH) can damage certain membranes and thus compromise their integrity.
• , ~
MOST PROBABLE NUMBER (MPN)ASSAYS
Principle Assessment of lytic viral activity requires that the virus particle destroys a host cell. Using a dilution approach, we can estimate the abundance of viruses in a sample. This process is based on the theoretical assumption that a single infectious virus can destroy a population of sensitive host cells (given time). The MPN approach to quantifying infectious viruses involves the exposure of a series of log-based dilutions of the sample containing the viruses to a liquid culture of host cells. Given an appropriate range of dilution (crossing the range where the mean viruses per aliquot sample is approximately 1.0) then the abundance of infectious viruses can be estimated. While the individual MPN exposures are scored '_+', comparison of these scores to MPN tables (or analysis by computer software) allows for an estimate of infective units. Replication can be achieved in several ways, with many labs now commonly using multi-well plates to enhance this (see Alternative technique). Ultimately, the desired levels of sensitivity and accuracy will dictate the volumes, number of replicates and scale of cultures to be used.
57
Equipment and reagents •
Culture medium for marine phytoplankton. ESAW (Harrison et al., 1980)
• • •
and its derivations commonly work well as a general growth medium. Liquid culture of host organism in exponential growth phase. 7 ml glass culture tubes (13 × 100 mm) with polypropylene screw caps. Fluorometer with filter set for chlorophyll determinations (Ks, 420ox;
•
>6400m). 25 mm, 0.22 lum nominal pore-size low protein binding filters (e.g.
•
Durapores®). Filtration funnel and receiver or Swinnex ® filter holder (and 10 cm 3
• •
syringe) for 25 mm filters. Pipette ( I - 5 ml) and tips for liquid dispensing. Culture facilities for phytoplankton.
Alternative technique (requires the following materials) • 96-well microtiter plates with lids. • Multichannel pipette and tips. • Fluorescent plate reader with filters for chlorophyll (Xs, 4200x; > 640om).
Application
(using 7 ml culture
tubes)
Collect water samples for screening into sterile p o l y p r o p y l e n e or polycarbonate containers and maintain them at 4°C in the dark until they are screened. Screen samples as soon as possible. Prior to screening, filter 25 ml of sample through a 0.22 p m nominal pore-size filter to remove bacteria, algae, protists, etc. In some cases other pore-size filters can be used (see Troubleshooting). From this, make a set of serial dilutions (10fold dilutions with sterile culture medium) with the sample to provide a dilution range of I to 10 4of the sample. A d d 1 ml of each of these dilutions to the exponentially growing host cultures (below) to screen for lytic activity. To prepare hosts for screening, transfer an aliquot of the host to fresh culture medium. For example, transfer 50 ml of an exponentially growing batch culture into 450 ml of medium. Monitor growth in this culture so that cells can be used as soon as the exponential phase of growth begins. As exponential growth begins, transfer 5 ml to each of fifty-five 7 ml culture tubes, assuming that ten replicates (and five controls with no sample added) is the desired n u m b e r for the experiment, and that five dilutions are being used (Suttle, 1993). Gently mix the tubes, record the fluorescence and place the tubes in culture facilities. Remove the tubes daily (for u p to seven days) and repeat the measurement of fluorescence. Cultures not clearing in seven days are assumed to not contain virus. For each dilution, record the n u m b e r of tubes that have cleared and use this data to calculate the MPN for the concentration of viruses in the sample. The MPN can be determined from published values in tables
58
(Koch, 1981) or by using c o m p u t e r programs which can also provide confidence intervals and standard estimates of error (Hurley and Roscoe 1983). Sample results are s h o w n in Figure 4.1.
500
400
Cyanophage P6 ! Cyanophage P56 [ Control !
T
i i
TJ soo t~ 0
T
200
m
ii 100
1
2
3
Time (days) Figure 4.1. Typical chlorophyll fluorescence from the cyanobacteria SyJlechococcus sp. WH7803 in culture with and without added viruses. The addition of cyanophage P6 demonstrates tile typical clearing seen in tubes during MPN assays. Both the control and cyanophage P56 (which does not infect this Synechococcus) demonstrate no clearing, and this was consistent up to 7 days (not shown).
Alternative technique As described by Suttle and Chan (1993) and Bratbak et al. (1998), microtiter plates can be substituted for culture tubes to screen cultures that will grow in these systems. Repeat the process as above, but substitute a 96-well microtiter plate for the 7 m l culture tubes and adjust volumes of host (100 tJl) and virus (50 1_ll). Maintain cultures u n d e r standard growth conditions and screen them daily, either visually or with a fluorescence plate reader equipped to monitor chlorophyll fluorescence. When using microtiter plates, it is easy to expand the dilutions from 5 to 7 (or more) tenfold steps. However, in the microtiter assay less sample is screened, so the m i n i m u m detection limits (sensitivity) of the assay is decreased.
Troubleshooting One problem c o m m o n l y associated with the screening of natural samples is the breakthrough of u n w a n t e d organisms (e.g. bacteria and protozoans) through the filter into the sample to be screened. This problem is often difficult to diagnose until after the experiment has been carried out, but samples can be examined by epifluorescence microscopy to determine the $9
presence of u n w a n t e d organisms if this problem becomes of concern. Another problem that occurs is the destruction or removal of viruses in the sample during storage, often by bacteria or protozoan grazers. To avoid this problem, filter samples upon collection (as described) and store in the dark at 4°C until use. It should be pointed out that viral infectivity will decay, even under these conditions. However, in at least one case infective viruses have been found in these concentrates after storage for 7 years under the above conditions (Wilhelm and Suttle, unpublished data). Another problem to consider is the removal of infectious viral particles by filtration. Different size filters are commonly suggested in different protocols (e.g. 1.0 lain, Suttle, 1993; 0.45 lain, Garza and Suttle, 1998). While we have suggested the use of a 0.22 lain filter in this protocol, it should be considered that decreasing the pore-size of the prefilter increases the possibility of viral retention during that step. In any case, all filters will retain some degree of viruses during this step, so consistency in pore-size, filter matrix and technique (e.g. pressure) is critical in providing reproducible results. Growth in the controls must also be closely monitored. Should all or a subset of the controls not grow, then it is not possible to determine if clearing in the test cultures is due to viral activity or non-experimental effects. It is therefore important to have established the ability to consistently grow the host culture in the lab prior to attempting to quantify viral particles. Finally, while the choice of using culture tubes relative to microtiter plates is left to the investigator, we would like to point out that the use of microtiter plates decreases the detection limit for infective viruses in the samples. As described above, the tube method will increase the detection limit 20-fold relative to the microtiter method (as 20 times more sample is screened).
,,,,~
P L A Q U E ASSAYS
Principle Plaque assays are commonly used in bacteriophage studies in order to enumerate the abundance of infectious phage in a sample. These same techniques m a y be applied to the enumeration of phycoviruses. Plaque assays have the advantage over MPN assays of providing increased accuracy, but have the disadvantage of requiring that hosts cells must be culturable and provide a confluent lawn on agar solidified growth medium. The principle of the plaque assay is simple: it assumes that, within a complete lawn of organisms on a Petri plate, each individual virus will produce a clearing or 'plaque' where it has lysed the localized host cell population. The plaque assay also provides the added advantage of allowing individual plaques to be isolated directly from the plate, providing a clonal copy of each virus. Moreover, in some cases the presence of turbid plaques can be taken as an indication of a potential lysogenic virus (although significant testing is required to confirm this). 60
Equipment and reagents • Culture medium for marine phytoplankton. ESAW (Harrison et al., 1980) and its derivations commonly work well as a general growth medium. • Agar for the solidification of culture medium (e.g. BactoAgar from Difco). • Liquid culture of host organism in exponential growth phase. • Autoclave/microwave. • Temperature controlled water bath or dry block. • Microfuge. • Vortex mixer. • 25 ram, 0.22 pm nominal pore-size low protein binding filters (e.g.
Durapores).
m
< O
• Petri places (plastic, 15 × 100 mm). • I and 5 ml pipeccers and tips. • 1.5 ml microfuge tubes. • 13 × 100 mm disposable borosilicate glass cubes and rack(s). • Erlenmeyer culture flask (250 ml). • Culture facilities for algae.
Application Prior to screening cultures, plates for the establishment of confluent lawns must be created. Bottom agar for these plates is created by adding 1~ ( w / v ) agar to the appropriate culture m e d i u m and autoclave sterilizing the sample. After the m e d i u m is allowed to cool to 60°C, pour plates (15 to 20 ml) u n d e r sterile conditions and allow them to solidify. It is important that plates are only p o u r e d to = 50% capacity. Once dried, invert the plates and store them as other cultures plates (4°C, dark). They are usually good for up to a week or more. Top agar is also required for plaque assays. To prepare it, add 0.6% ( w / v ) agar to 100 ml of growth m e d i u m in a 250 ml flask or media bottle. If sealed after sterilization, this can be stored at room temperature. When required, the agar is remelted in a microwave oven and 2.5 ml aliquots placed into three 7 ml disposable culture tubes per sample to be screened. Maintain these tubes at a temperature between 45 and 47°C in the water b a t h / h o t block until use. Fill three tubes with top agar to use as controls. To prepare water samples for screenings, filter 25 ml of sample through a 0.22 lJm pore-size filter to remove bacteria, algae, protists, etc. as these organisms m a y cause false plaques to form. It might be necessary with some samples to carry out a series of dilutions prior to undertaking the plaque assay, as it is desirable to have only 20-200 infectious viruses per aliquot. These dilutions can be carried out as described above, using sterile marine m e d i u m and a series of culture tubes. To begin the plaque assay, start cultures of phototrophs to allow for yields of around 10 r ml ' of exponentially growing cells. Harvest the cells by gentle centrifugation (3000-5000g) and then resuspend them to around 1if' cells ml ~. When working with heterotrophic bacteria this 61
4.1
¢-
concentration step is generally not required. Transfer cells (500 lad to three sterile microfuge tubes, and 500 lJl of sample to each tube, then close the tubes and mix by vortexing. For each set of experiments add 500 ~1 of sterile culture m e d i u m to hosts in microfuge tubes to act as a control. After samples are combined in microfuge tubes, a brief spin in a microfuge will remove any liquid from the interior lid of the tubes. Allow samples to sit so that the virus can adsorb to the host cells. Adsorption times for heterotrophic bacteria c o m m o n l y range from 5-25 minutes, but 30-45 minutes is sufficient w h e n the kinetics of adsorption are u n k n o w n (Suttle, 1993). After the adsorption period, mix the contents of each microfuge tube with a tube of top agar by vortexing. Quickly p o u r this mixture onto the bottom agar in the Petri plate, and 'swirl' the sample on a flat surface to evenly distribute the top agar mixture. After the plates dry and solidify (=60 minutes), invert them and m o v e them to appropriate culture facilities. Enumeration of plaques on the plates occurs once confluent host lawns have established (2-6 days). Individual plaques are e n u m e r a t e d and assumed to represent the presence of one lytic virus in the samples. For statistical relevance, it is desirable to enumerate plates from dilutions with plaque abundances ranging from 20-200 per plate. Once plaques are enumerated, the abundance of viruses infecting the host can be determined as follows: A = (p / d) x ( 1 / v ) where A is the abundance of infectious viruses (ml '), p is the n u m b e r of plaques on the plate, d the dilution factor for that plate and v the v o l u m e (ml) of sample a d d e d (as described above, 0.5). A comparison of the two assays (most probable n u m b e r and plaque) is given in Table 4.1.
Table 4.1 Comparison of the most probable number assay with the plaque assay for the enumeration of phycoviruses Method
Advantages
Disadvantages
MPN assay
• Flexible with culture requirements • Amenable to high replication • Does not require growth on solidified medium • Enumerates only infective viruses • Higher accuracy than MPN assay • Provides indication of potential lysogens • Provides for easy purification of viruses • Enumerates only infective viruses
• Less precision than plaque assay • Low probability of detecting lysogens
Plaque assay
62
• Must be cultured on solidified medium • Natural bacteria can cause plaqueqike clearings
Troubleshooting The most significant problem associated with variations in plaque assay results is inconsistency of technique. Ensuring that cells are in the same phase of growth in each experiment is vital to providing reproducible results. As well, culture conditions (including temperature) should be held constant to allow for an intercomparison of samples. To account for variation, positive controls consisting of samples with a k n o w n abundance of infectious viruses can be included in every experiment. However, it should be r e m e m b e r that stored samples of viruses slowly lose infectivity over time, so samples that are examined 6 m o n t h s apart may not be comparable using the same positive control. It should be stressed that the plaque assay m e t h o d is often difficult to use with eukaryotic plankton (the exception being strains of Chlorella spp.). As well, certain cyanobacteria will not grow on standard agar, as it usually contains too m a n y impurities. Better growth of cyanobacteria on agar plates can be achieved by first removing these impurities (Waterbury and Wiley, 1988).
CONCLUSIONS Two methods for the enumeration of infectious phycoviruses as well as a m e t h o d to increase the concentration of viruses in a water sample are discussed here. Each m e t h o d for viral enumeration has its particular advantages and disadvantages (Table 4.1). The choice of the particular m e t h o d will ultimately d e p e n d on the culturability of the host ceils in question.
References Bratbak, G., Egge, J. and Heldal, M. (1993). Viral mortality of the marine alga Emiliania huxleyi (Haptophyceae) and termination of algal blooms. Mar. Ecol. Prog. Ser. 93, 39 48. Bratbak, G., Jacobsen, A., Heldal, M., Nagasaki, K. and Thingstad, E (1998). Virus production in Phaeocystis pouchetii and its relation to host cell growth and nutrition. Aquatic Microbial Ecol. 16, 1 9. Chen, E and Suttle, C. A. (1996). Amplification of DNA polymerase gene fragments from viruses infecting microalgae. Appl. EHvipx~n.Microbiol. 61, 1274-1278. Chen, E, Suttle, C. A. and Short, S. M. (1996). Genetic diversity in marine algal virus communities as revealed by sequence analysis of DNA polymerase genes. Appl. Environ. Microbiol. 62, 2869-2874. Cottrell, M. T. and Suttle, C. A. (1995). Dynamics of a lyric virus infecting the photosynthetic marine picoflagellate Micromonas pusilla. LiutHol. Oceanog. 40, 730 739. Field, C., Behrenfeld, M., Randerson, J. and Falkowski, P. (1988). Primary production of the biosphere: integrating terrestrial and oceanic components. Science 281, 237-240. Fuhrman, J. A. (1999). Marine viruses and their biogeochemical and ecological effects. Nature 399, 541-548.
63
Fuller, N. J., Wilson, W. H., Joint, 1. R. and Mann, N. H. (1998). Occurrence of a sequence in marine cyanophages similar to that of T4 g20 and its application to PCR-based detection and quantification techniques. Appl. Environ. Microbiol. 64, 2051-2060. Garza, D. R. and Suttle, C. A. (1998). The effect of cyanophages on the mortality of Synechococcus spp. and selection for UV resistant viral communities. Microbial Ecol. 36, 281-292. Harrison, P., Waters, R. and Taylor, E (1980). A broad spectrum artificial seawater m e d i u m for coastal and open ocean phytoplankton. J. Phycol. 16, 28-35. Hurley, M. and Roscoe, M. E. (1983). Automated statistical analysis of microbial enumeration by dilution series. J. Appl. Bacteriol. 55, 159-164. Jocobsen, A., Bratbak, G. and Heldal, M. (1996). Isolation and caracterization of a virus infecting Phaeocystis pouchetii (Prymnesiophyceae). J. Phycol. 32, 923 927. Koch, A. L. (1981). Growth measurement. In: Mamml of Methods for General Bacteriology (P. Gerhardt, Ed.), p. 179. American Society of Microbiologists, Washington, DC. Lawrence, J. E., Chan, A. M. and Suttle, C. A. (2000). A novel virus causes lysis of the toxic bloom-forming alga, Heterosigma akashiwo (Raphidophyceae). J. Phycol. (in press). Milligan, K. L. D. and Cosper, E. M. (1994). Isolation of virus capable of lysing the brown tide microalga, Aureococcus aJiophagt~fferens. Science 266, 805-807. Nagasaki, K. and Yamaguchi, M. (1997). Isolation of a virus infectious to the harmful bloom causing microalga Heterosi~ma akashiwo (Raphidophyceae). Aquatic Microbial Ecol. 13, 135 140. Rodda, K., Clark, L., IngaI1, E. and Suttle, C.A. (1996). Infective cyanophages persist in anoxic sediments on the continental shelf of the Gulf of Mexico. EOS Trans., Am. Geophys. UuioJl 76(3), OS 51 I-6. Short, S. M. and Suttle, C. A. (1999) Use of the polymerase chain reaction and denaturing gradient gel electrophoresis to study diversity in natural virus communities. Hydrobiolosqa 401, 19-32. Suttle, C. A. (1993) Enumeration and isolation of viruses. In: Handbook of Aquatic Microbial Ecolok,y (P. E Kemp, B. E Sherr, B. B. Sherr, and J. J. Cole, Eds), pp. 121-134. Lewis Publishers, Ann Arbor, MI. Suttle, C. A. (1996). Community structure: viruses. In: Manual of Environmental Microbiology (C. Hurst, G. Knudson, M. Mclnerney, L. Stezenbach and M. Walter, Eds), pp. 272-277. ASM Press, Washington DC. Suttle, C. A. and Chan, A. M. (1993). Marine cyanophages infecting oceanic and coastal strains of SyHechococcus - - abundance, morphology, cross-infectivity and growth characteristics. Mar. Ecol. ProS. Set. 92, 99-109. Suttle, C. A. and Chan, A. M. (1994). Dynamics and distribution of cyanophages and their effect on marine Synechococcus spp. Appl. Environ. Microbiol. 60, 3167-3174. Suttle, C. A. and Chan, A. M. (1995). Viruses infecting the marine Prymnesiophyte Chrysochromulina spp.: isolation, preliminary characterization and natural abundance. Mar. Ecol. Pro,~¢.Ser. 118, 275-282. Suttle, C. A., Chan, A. M. and Cottrell, M. T. (1991). Use of ultrafiltration to isolate viruses from seawater which are pathogens of marine phytoplankton. Appl. E~lviron. Microbiol. 57, 721-726. Waterbury, J. B. and Valois, E W. (1993). Resistance to co-occurring phages enables marine Synechococcus communities to coexist with cyanophages abundant in seawater. Appl. Environ. Microbiol. 59, 3393-3399. Waterbury, J. B. and Willey, J. M. (1988). Isolation and growth of marine planktonic cyanobacteria. Meth. Enzymol. 167, 100-105.
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Wilhelm, S. W. and Suttle, C. A. (1999). Viruses and nutrient cycles in the sea. BioScience 49, 781-788. Wommack, K. E., Hill, R. T. and Colwell, R. R. (1995). A simple method for the concentration of viruses from natural water samples. J. Microbiol. Meth. 22, 57-67.
List of suppliers The following is a selection of companies. For m o s t products, alternative suppliers are available.
Fisher Scientific Worldwide 50 Fadenl Road Springfield NJ 07081-3193, USA Phone: 1 973 467- 6400 Fax: I 973 376-1546 Disposables, m e d i a reagents, vortexers, microfuges
Millipore 80 Ashby Road Bedford, M A 01730, USA Phone: (800) MILLIPORE Fax: 1 781 533-3110 Ultrafiltration systems, m e m b r a n e filters a n d holders
6S
Turner Designs, Inc. 845 W. Maude Avenue Sunnyvale CA 94086, USA Phone: 1 408 749-0994 Fax: 1 408 749-0998 http://www.turuerdesis,,ns.conl Fluorometers
5 Estimating Viral Proliferation in Aquatic Samples Rachel T Noble' and Grieg F Steward 2 ~Southern California CoastalWater Research Project (SCCWRP),Westminster, CA USA;2Monterey Bay Aquarium Research Institute, Moss Landing, CA USA
CONTENTS
Introduction Bacteriophage production by 3H-rhymidine incorporation Calculation from bacterial productivity, the fraction of infected bacteria, and burst size Tracer dilution using fluorescently labeled viruses (FLV) Conclusions
~,4,eeee INTRODUCTION It is only within the last decade that m a r i n e viruses were d e t e r m i n e d to be consistently the m o s t a b u n d a n t biological entities in the sea (Fuhrman, 1999). Since then, m a n y a d v a n c e s h a v e been m a d e in u n d e r s t a n d i n g viral ecology (Fuhrman, 1999; Wilhelm and Suttle, 1999). Initial discoveries s h o w e d that viruses are a b u n d a n t in the ocean and that m a n y bacteria are infected with viruses (Bergh et al., 1989; Proctor and F u h r m a n , 1990). These data led researchers to believe that viruses are an i m p o r t a n t source of mortality in m a r i n e microbial food webs, but only p r o v i d e d a static picture. Subsequent studies have s h o w n that virus p o p u l a t i o n s are extremely dynamic, and can change quickly over short timescales (Bratbak et al., 1990; 1996). Estimates of viral production and decay rates p r o v i d e d the valuable confirmation that viruses are active m e m b e r s of the m a r i n e c o m m u n i t y (Heldal and Bratbak, 1991; Steward et al., 1992b). Viral p r o d u c t i o n involves the lysis of host cells and the release of cellular material as dissolved and colloidal organic carbon. Therefore, m e a s u r e m e n t s of viral replication rates are also useful for assessing the contribution of viruses to bacterial mortality and organic matter cycling in the ocean. By a s s u m i n g a burst size, viral productivity can be used to estimate rates of bacterial lysis. This a p p r o a c h provides an additional m e a n s to assess bacterial mortality along with the visualization of intracellular viral particles b y transmission electron m i c r o s c o p y (Proctor and F u h r m a n , 1990). Accurate m e a s u r e m e n t s of viral p r o d u c t i v i t y and t u r n o v e r are required to METHODS IN MICROBIOLOGY,VOLUME 30 ISBN 0 12 521530-4
Copyrigh{ © 2001 Academic Press Ltd All rights of reproduction in any torm reserved
properly model viral dynamics and their impacts upon aquatic microbial food webs. So far, however, there is no standard method for measuring viral productivity. A wide variety of different approaches have been used each with associated advantages and disadvantages. These methods include: 1. quantifying net increases in viral abundance over time (Bratbak et al., 1990); 2. measuring rates of viral decay (Heldal and Bratbak, 1991); 3. estimating viral DNA synthesis rates by radiolabeling (Steward et al., 1992a,b); 4. calculating expected viral release rates from estimated rates of bacterial lysis and an assumed burst size (Weinbauer et al., 1993); 5. measuring tracer dilution rates using fluorescently labeled viruses (FLV) as tracers (Noble and Fuhrman, 2000). The first approach, observing net increases in viruses over time, is the simplest means of demonstrating viral proliferation. However, use of this method is restricted to times when viral abundance is increasing and only provides a minimum estimate of productivity unless the viral decay rate is also known. The productivity estimates obtained are also dramatically influenced by the timescale of sampling (Bratbak et al., 1994, 1996). In the second approach, virus production is prevented by poisoning or removing host organisms and the rate at which viruses disappear (or decay) is observed. If the system was initially in steady state, the decay rate is assumed equal to the original rate of production. These two approaches have been used in a number of studies and variations on them are possible. Due to space limitations, however, this chapter focuses only on the last three methods listed above.
4'4'4'e~l'4l' B A C T E R I O P H A G E P R O D U C T I O N 3H-THYMIDINE INCORPORATION
BY
Principle Tritiated thymidine (~H-TdR) is taken up by bacteria, resulting in a radiolabeled intracellular nucleotide pool. During viral replication, :~H-TdR is drawn from the host's internal pools and incorporated into viral DNA. After labeling, a series of steps are followed to ensure specific measurement of ~H-TdR in virus-like DNA. First, viruses are separated from other organisms by 0.2 IJm filtration. Then, in the filtrate, putative viral DNA is distinguished from dissolved DNA by nuclease digestion (the DNA of intact viruses is protected from digestion). Finally, hydrolysis by hot acid or hot base is used to distinguish ~H incorporated in DNA from nonspecific incorporation that can result from metabolism of -~H-TdR by the cell. After the hydrolysis, remaining macromolecules are precipitated with cold trichloroacetic acid (TCA) and washed free of hydrolysis products. To minimize quenching, pellets are hydrolyzed with hot acid, then 68
the incorporated ~H is quantified by liquid scintillation counting. The rate of ~H-TdR incorporation is converted to viral productivity using an empirical conversion factor.
Equipment and reagents
• • • • •
• • • • • • • • • • • •
Polycarbonate tubes or bottles with screw caps (30 ml capacity) for sample incubations Pipets, micropipettors and tips Syringes (5 ml capacity) 0.2 lum syringe-tip filters (Acrodisc, Gelman) Plastic, sterile tubes for collecting filtrate (5 ml capacity) and for nuclease digestions (2 ml capacity) Microcentrifuge tubes (2 ml capacity) with screw-caps and o-rings: if following Alternative 2 (below) the tubes should have separate lids. For tubes that come with attached caps, the connector between tube and cap can interfere with placing the tube in a scintillation vial. Screw caps with orings are recommended to prevent leakage of radioactive material during centrifugation Vortex mixer *Filtration manifold for 25 mm filters *Mixed cellulose acetate and nitrate filters (25 mm, Millipore HAWP) and filter forceps tMicrocentrifuge Hot water baths Scintillation vials (7 ml capacity) Liquid scintillation counter Protective clothing, containers and facilities for handling radioactivity, and for collection and disposal of radioactive waste [methyl-3H]-thymidine (3H-TdR); specific activity ca. 3 TBq mmol ' Nuclease cocktail (contains deoxyribonuclease I and ribonuclease A, each at I U lal ', and micrococcal nuclease at 5 U IJl ') Formalin (37% w:v Formaldehyde solution) Trichloroacetic acid (TCA). Prepare working solutions of 70, 20 and 5% T C A in water (w:v). A 100%TCA stock solution is made by adding 227 ml of water to 500 g of TCA NaOH (5 M)
• • Scintillation cocktail
*Only required if following Altcrtu~tivc I (below) 'Only required if following AlterJu~tivc 2 (below)
Assay 1. A d d ~H-TdR (10 nM final conc., ca. 30 kBq ml ~) to duplicate 25 ml seawater samples and incubate at in situ temperature. 2. Collect a 4.5 ml subsample from each replicate at each of four or five time points taken between 0 and 18 h.
69
.
Filter the subsample through a 0.2 p m syringe-tip filter.
4. Split the filtrate into duplicate 2 ml samples. 5. A d d 10 1.11of nuclease cocktail to each and incubate at ca. 20°C for
lh. 6. A d d formalin to preserve the sample and store at 0 to 4°C until all time points have been collected. 7. Split each tube into duplicate 900 pl aliquots. 8. Add 100 pl of 5 M N a O H to one replicate and 300 ~1 of 20% TCA to the other and mix by brief vortexing. 9. Incubate one hour at 60°C (NaOH treatment) and the other at 90°C (the TCA-treatment). 10. Chill the tubes on ice for 5 rain. 11. Add 200 pl of cold 70% TCA to the NaOH-treated sample only, mix by brief vortexing, and incubate both tubes on ice an additional 10 min. Alternative I
12. Shake tubes to resuspend any precipitate and filter through 0.2 p m HAWP Millipore filters. 13. Rinse each tube with 1 ml 5% TCA and pass through the same filters. 14. Rinse filters three times with a few milliliters of cold 5% TCA. 15. Place filters in scintillation vials and add 0.5 ml 5% TCA. 16. Incubate at 90°C for 30 min then cool to RT. 17. A d d 5 ml of scintillation cocktail and vortex mix. 18. Count by liquid scintillation with quench correction. Alternative 2
12. Centrifuge tubes at 16 000g at 4°C for 10 min in refrigerated microcentrifuge. 13. Aspirate supernatant. 14. A d d 1 ml ice-cold, 5% TCA then cap and invert tube to rinse sides. 15. Centrifuge as before for 5 min. 16. Aspirate supernatant. 17. Add 50 t~1 of 5% TCA, mix by vortexing, then heat to 90°C for 30 min. 18. Add 1.4 ml scintillation cocktail and vortex mix. 19. Place tube in a 7 ml capacity plastic scintillation vial. 20. Count by liquid scintillation with quench correction.
Calculations The rate of incorporation is derived from a linear regression of d p m vs. time. If there is an initial lag in incorporation, the T,, time point is eliminated and regression is p e r f o r m e d on the remaining points. An initial lag in incorporation might be expected for two reasons: (1) it would reflect the 70
portion of the latent period between cessation of DNA synthesis and release of intact phage; and (2) if viruses recycle DNA then the first viruses released w o u l d have a v e r y low specific activity. The d p m rate is converted to a TdR incorporation rate based on the specific activity of the a d d e d 3H-TdR as reported by the manufacturer. The TdR incorporation rate is then converted to a viral production rate using an empirically determined conversion factor that accounts for intracellular isotope dilution. The available estimates of conversion factors suggest a value of about 102~ viruses per mole TdR incorporated.
T h e conversion factor The empirical conversion factor is obtained from incubation experiments in which both ~H-TdR incorporation and viral abundance are followed with time. This approach requires a non-steady state system, i.e. one in which viral a b u n d a n c e is increasing. Viral abundance is plotted vs. moles ~H-TdR incorporated and the conversion factor is taken as the m a x i m u m slope of the curve. The time points used in the curve should be within the range of those used for measuring productivity (i.e. < 24 h). The accuracy of the conversion factor d e p e n d s on the degree to which a net change in viral a b u n d a n c e reflects gross viral production. Usually there will be some simultaneous decay of viruses that will tend to result in underestimation of the factor. Manipulation of a seawater sample m a y be useful for inducing a non-steady state. For example, a seawater culture approach was e m p l o y e d by Steward et al. (1992a). Empirical estimates of the conversion factor from this type of m i x e d - c o m m u n i t y approach m a y reveal useful information which w o u l d not be revealed with the use of pure culture studies. In this case, differential centrifugation of seawater was used to prepare an inoculum containing bacteria and viruses (the supernatant after centrifuging at low speed to remove particles and larger cells) and a virus-free diluent (the supernatant after ultracentrifugation). The inoculum was diluted 50-fold into the virus-free water and split into parallel samples to follow virus counts and ~H-TdR incorporation. Reducing the background pool of viruses resulted in a non-steady state system in which viral production temporarily exceeded decay. Since a conversion factor is not trivial to obtain, all studies of virus production by radiolabeling so far have relied on the conversion factors derived by Steward et al. (1992a). The conversion factor 2 x 102' viruses per mole TdR was a d o p t e d in a study by Steward et al. (1996), but the more conservative factor of 6 x 102~'viruses per mole TdR was applied in other studies (Fuhrman and Noble, 1995; Kepner et al., 1998). Empirically derived conversion factors have been required due to our ignorance about some aspects of viral replication in natural marine assemblages. Two important u n k n o w n s are (1) the g e n o m e size of the replicating viruses and (2) the m a g n i t u d e of intracellular isotope dilution. Recent data suggest that the average g e n o m e size in marine viral assemblages is fairly consistent at ca. 50 kb (Steward and Azam, 2000; Steward et al., 2000). From this information we can make a theoretical estimate of a
71
conversion factor. If we assume the average viral genome to be 50 kb of dsDNA with an average GC content of 50%, then the theoretical conversion factor is 2.4 x 101. viruses per mole TdR. This factor is substantially lower than the empirical factors above, suggesting that isotope dilution is also important. Two potential causes of intracellular isotope dilution have been identified. One is dilution of the intracellular free nucleotide pool as a result synthesis by de novo or rescue pathways (Moriarty, 1986). A comparison with ~2PO~ incorporation suggested that the isotope dilution of ~H-TdR during bacterial DNA synthesis is about 5-fold (range 3 to 7; Fuhrman and Azam, 1982). If viruses draw from the same nucleotide pool as the bacteria, they would face the same isotope dilution. Multiplying the theoretical factor by five increases it to about 1.2 x 10~ viruses per mole TdR. This is still substantially lower than that observed, suggesting additional sources of isotope dilution. A second potential cause of isotope dilution is specific to measurements of viral DNA synthesis. Wikner et al. (1993) developed a mathematical model which showed that isotope dilution during viral DNA synthesis can vary dramatically depending on the source of the nucleotides. If a virus uses degraded host DNA as a source of nucleotides, the effective specific activity of the incorporated thymidine is drastically reduced. In this case, viruses draw from a nucleotide pool that equilibrates relatively slowly, and at a rate which depends on the growth rate of the host. The model was developed for ~-'P radiolabeling, but the dilution effect would be similar for 3H-TdR. Assuming an average growth rate for marine bacteria of 0.02 h ~,the model predicts that isotope dilution between 6 and 18 h would vary from 15- to 6-fold and at the midpoint (12 h) would be 8.3-fold. Combining this with the isotope dilution factor for ~H-TdR incorporation by bacteria (5-fold) results in total isotope dilution factor of about 42. Multiplying the theoretical factor by this total potential isotope dilution raises the theoretical factor to 1 x 10-", which is in the range of observed empirical factors. Although there is considerable uncertainty in the predicted isotope dilution, the reasonable correspondence between predicted and observed factors suggests that nucleotide recycling may be common among marine bacteriophages.
Applications The ~H-TdR incorporation method estimates only the production of DNA viruses infecting bacteria. Radiolabeled phosphate (~'PO~~) has also been used following a similar procedure (Steward et al., 1992a,b). Presumably this would include production of phytoplankton viruses as well. However, for measurements with ~2p (or ~P), the phosphate concentration of each sample must also be measured in order to account for extracellular isotope dilution. These isotopes also have shorter half-lives and are more challenging to handle. For these reasons only the ~H-TdR method has been presented here. The radiolabeling method has been used to estimate viral productivity in a variety of environments including coastal and offshore waters of the Southern California Bight (Steward et al., 1992b; 72
Fuhrman and Noble, 1995), in the Bering and Chukchi Seas (Steward et al., 1996) and in an oligotrophic Antarctic lake (Kepner et al., 1998). This method is a fairly simple extension of a routine bacterial productivity method (Fuhrman and Azam, 1982), requiring only a limited amount of extra equipment, and reagents that are fairly inexpensive. The method appears to be better suited for more productive waters (i.e. > 1 x 10" viruses 1 ~d ~). So far, significant rates in the ocean have been measurable only in surface waters (< 30 m depth). The major disadvantages of this approach are: (1) that it requires the use of a conversion factor that is still poorly constrained; and (2) that trapping of viruses (e.g. those adsorbed to particles) during 0.2 ~tm filtration m a y result in an underestimation of phage productivity.
C A L C U L A T I O N FROM BACTERIAL P R O D U C T I V I T Y , THE F R A C T I O N OF INFECTED BACTERIA, A N D BURST SIZE Principle During the late stages of viral infection, mature viruses are assembled within an infected bacterium. At this stage, infected bacteria are recognizable by transmission electron microscopy and are referred to as 'visibly infected'. The percentage of the total infection cycle during which a cell is visibly infected has been empirically determined and can be used to estimate the total number of infected cells in a sample (Proctor et aI., 1993). The fraction of bacterial mortality due to viruses can then be estimated with some further assumptions concerning the relationship between the virus latent period and the host generation time (Proctor et al., 1993; Binder, 1999). The fraction of mortality due to viruses is multiplied by the bacterial productivity in the sample to obtain a rate of cell lysis. Assuming a burst size, the rate of viral production can then be estimated.
Equipment and reagents Bacterial productivity • Equipment and reagents are as described in Chapter production'.
I1: 'Bacterial
Transmission electron microscopy • Ultracentrifuge and swinging-bucket rotor (SW41 or equivalent) • Ultracentrifuge tubes (with adapters to create a flat surface for grid support). Adapters can be molded using epoxy (Borsheim et al., 1990) or machined from plexiglass (Wells and Goldberg, 1992) • Pipets, micropipettors and tips • Glutaraldehyde, 25 or 50% stock, electron microscopy grade • Formvar-coated, 200 or 400-mesh copper grids
73
• •
•
Forceps, fine tip, self-closing Uranyl acetate, 0.5% in water. Uranyl acetate is toxic and radioactive so minimal amounts should be used. Using an analytical balance, as little as 5 to I0 mg can be weighed to make a I to 2 ml batch in a microcentrifuge tube. Stir or agitate for half an hour to dissolve Transmission electron microscope
Assay 1. 2. 3.
4.
5.
6.
Bacterial productivity is measured as described in Chapter 11: Bacterial Production. Aliquots from the same sample are fixed with glutaraldehyde (1% final conc., v:v). Bacteria are harvested from fixed seawater samples onto 400-mesh copper grids via ultracentrifugation at 60 000 to 80 000g for 20 min. The centrifugation speed and time are lower than those used to pellet viruses as this is thought to minimize disruption of infected cells (Weinbauer et al., 1993). Grids are removed and stained for 30s by dipping into 0.5% uranyl acetate, then rinsed three times for 10 s in aliquots of 0.02 micron filtered water. Bacteria are examined by electron microscopy at 100 kV accelerating voltage at a magnification ranging from x 30 000 to × 50 000. Bacteria are scored for the presence of intracellular viruses. Cells containing a m i n i m u m of five phages are considered infected and are p h o t o g r a p h e d for documentation and further examination of cells (Weinbauer and H6fle, 1998). Confidence intervals for the proportion of infected cells can be calculated from the relationship between the F distribution and the binomial distribution (Zar, 1996). The n u m b e r of cells to count will d e p e n d on how m u c h error is acceptable in the estimate. As a rule of thumb, count u p to 1000 total bacteria or ten infected bacteria, whichever n u m b e r is reached first. Photographs of infected cells can be examined to estimate the n u m b e r of viruses per cell (also k n o w n as 'burst size').
Calculations The fraction of visibly infected bacterial cells (FVIC) is converted to the actual fraction of infected cells (FIC) and then converted to the fraction of mortality due to viral lysis ( F M V L ) using the formulae presented by Binder (1999): FIC ~ 7.1 F V I C - 22.5 F V I C :
and F M V L -~ (FIC + 0.6FIC 2) / (1 - 1.2FIC)
74
Viral productivity (VP) is then calculated as: V P ~ F M V L x BP x N ,
where BP is bacterial productivity (in units of cells 1 ' d ~), and N, is the n u m b e r of viruses released per lysed cell (i.e. burst size, in units of viruses per cell). In the ideal case, burst sizes could be estimated for each experiment by examining photographs of infected cells. Preferably, only cells completely full of viruses w o u l d be included in the burst size calculation, but this could severely limit the n u m b e r of usable cells. In addition, burst sizes are still likely to be underestimated due to some viruses being obscured by overlying ones. A burst size based on only a few cells in a sample could be severely biased so in m a n y cases it m a y be best to assume a range of burst sizes and apply the same range to all samples. TEM-based estimates of burst sizes cited in the literature range from 6 to 300 for individual cells, but m e a n values are typically a r o u n d 20 to 50.
Applications This m e t h o d has been used in both marine (Weinbauer et al., 1993; GuixaBoixereu et al., 1996; Steward et al., 1996) and freshwater (Hennes and Simon, 1995; Weinbauer and H6fle, 1998) environments. An attractive feature of the m e t h o d is that it provides a detailed view of viral infections in the bacterial community. The incidence of infection and burst sizes can be d e t e r m i n e d separately for bacteria of different morphologies (Weinbauer and Peduzzi, 1994). An advantage of this m e t h o d is that, in situations where bacterial productivity is already being measured in the field, little extra effort is required to obtain samples for viral productivity estimates. The subsequent laboratory analysis, however, is s o m e w h a t time consuming, tedious and expensive as it can require about an hour of continuous TEM beam time per sample. The most significant disadvantage of the m e t h o d is the great deal of uncertainty in a n u m b e r of the factors used to derive the equations. In particular, it is assumed that (1) infected and uninfected bacteria are grazed equally, (2) latent period equals bacterial generation time, and (3) that, on average, viruses are only visible during the last 18.4% of the latent period. The validity of these assumptions in natural ecosystems remains unknown. In addition, burst sizes for natural communities are difficult to obtain with certainty (Weinbauer et al., 1994).
******
TRACER DILUTION USING FLUORESCENTLY L A B E L E D VIRUSES (FLV)
Principle Recent studies have d e m o n s t r a t e d the use of epifluorescence microscopy paired with fluorescent stains like DAPI, SYBR Green I, and Yo-Pro I for counting virus particles (Hara et al., 1991; Hennes and Suttle, 1995; 75
Weinbauer and Suttle, 1997; Xenopolous and Bird, 1997; Noble and Fuhrman, 1998). This method utilizes SYBR Green I stained FLVs as tracers for determining rates of viral production and removal. This m e t h o d was adapted from, and is mathematically similar to the 'isotope dilution m e t h o d ' used to measure the release and uptake of amino acids with radioisotopes. The isotope dilution technique was described previously by Blackburn (1979), Fuhrman (1987), and Glibert (1982), but will not be described in detail here. Basically, the FLV are analogous to labeled molecules used as tracers of amino acids in earlier studies. When FLV are added into the seawater at tracer levels, two counts are made: one count is performed on an unstained sample to enumerate only the FLV, and then a second count for the total virus abundance. Virus removal processes decrease the n u m b e r of FLV and unstained viruses proportionly, because the FLV act as a tracer. However, virus production produces only unlabeled viruses, thereby diluting the initial pool of FLV. Using the rate of change of both labeled and unlabeled viruses over time, virus production and removal rates can be calculated.
Equipment and reagents
Counts of viruses and FLV • Required equipment and reagents for epifluorescence counts of viruses are outlined in Chapter 3. Enumeration of viruses, and are also detailed in, Noble and Fuhrman (I 998). Preparation of FLV • 20 I low density polyethylene Cubitainers • 142 mm stainless steel filtration units (Advantec MFS, Inc.) • 20 or 40 I stainless steel pressurized container (Advantec MFS, Inc.) • 142 mm glass fiber filters (Gelman Sciences, Inc.) • 142 mm Durapore filters (0.22 ~tm, Millipore, Inc.) • Tangential flow ultrafiltration system with a 30 kD MW cutoff spiralwound membrane cartridge (SLY30, Millipore, Inc.) • Centriprep-30 centrifugal ultrafiltration units (Millipore, Inc.) • Whirl-pak bags Assay
Preparation of FLV tracers 1.
Filter 20 1 of seawater at 5 kPa serially through a 142 m m diameter glass fiber filter and a 0.22 p m pore-size Durapore filter to remove bacteria and protists. Make sure to use non-toxic tubing for the filtration. Note: for smaller-scale experiments, it is possible to concentrate a smaller v o l u m e of seawater to make the virus concentrate. Prefilter with 47 m m diameter glass fiber and 0.2 Hm filters, and concentrate the filtrate with either Midgee H o o p s (AG Technology, Inc.) or Centriprep-30 units to a final v o l u m e of 5 to 10 ml. This is especially useful on-board ship.
76
.
Concentrate viruses in the filtrate to ca. 150 ml by tangential flow ultrafiltration. Perform further concentration using Centriprep-30 ultrafiltration units to a final v o l u m e of ca. 5 ml.
Perform the following steps involving the SYBR Green I (Molecular Probes, Inc.) stain in dark or s u b d u e d light. 3.
4.
5.
6.
Transfer the virus concentrates to new Centriprep-30 units, and add the SYBR Green I at a final concentration of 2.5%. Incubate in the dark for at least 8 hours at 4°C. After the staining period, spin the FLV concentrate at 3000g for 15 min to 2-3 ml. To rinse away u n b o u n d stain, add 5 ml of 0.02 gm filtered seawater to the concentrate and reconcentrate to 2-3 ml. Repeat the rinse twice more. Recover the final concentrates in a total of 5 ml of 0.02 g m filtered seawater. If the concentrate is not to be used immediately, it can be stored at -20°C for several months. Immediately dilute 10 ~1 of the concentrate to a final v o l u m e of 2 ml with 0.02 g m filtered seawater. FLV concentration is determined by epifluorescence microscopy following the same procedure as described for total counts of viruses by SYBR Green I (Chapter 3) except that the filters are not stained after filtering. At the same time, determine the ambient virus concentration in the original seawater sample by following the usual SYBR Green l m e t h o d (Chapter 3: Enumeration of viruses).
Experimental p~tocol 1.
2.
3. 4.
5.
A d d 350 ml of seawater to each of two Whirl-Pak bags (if possible, replicate these treatments). To one bag, add 7 ml of 0.02 ~m filtered formalin (final concentration, 2 ~). Working in s u b d u e d light, add FLV concentrate to each of the seawater samples at tracer levels of viruses (<10% of the original virus concentration). Incubate the seawater samples in the WhirlPak bags in the dark at ambient seawater temperatures for 6-16 hours, with 40 ml subsamples taken at time zero and every few hours. Optimally, experiments should be started in the evening, thereby simulating in situ production and removal rates during the nighttime period of incubation. Fix each subsample immediately after collecting by adding I ml of 0.02 lam filtered formalin. Determine the total virus concentration by the SYBR Green I staining m e t h o d (Chapter 3), and the FLV concentration by the same procedure without the staining step. The volumes required are typically 1 to 2 ml for total virus counts and 10 ml for FLV counts. Slides should be counted immediately but can be stored in the dark at -20 ° C. Count a total of > 200 FLV per filter.
77
Calculations Production and r e m o v a l rates are calculated using equations a d a p t e d from Glibert (1982) and F u h r m a n (1987), w h e r e the input variables are incubation time (t), total virus concentration at time zero (C,,) and at time t (C,), and FLV concentration at time zero ( F L V , ) and at time t (FLV,). The decay constant, k, is calculated as:
k-
In (R o / R t )
w h e r e RL,and R, are the ratios of F L V to total viruses at time zero and time t, respectively. For example, R, is FLV,, divided by C~,. The m e a n specific activity, R (analogous to specific activity in radioisotopes), is then calculated as
R
The viral decay or r e m o v a l rate, D,,, is calculated as, D v = (FCVo - F L V t )
(Rb~~ x t) and the viral p r o d u c t i o n rate, P,, is calculated as
PF =
In ( R o / R t ) in (C,, / C t ) x t
x (C o - G )
Troubleshooting One of the m a i n sources of viral loss in the formalin-treated samples is fading of the viruses over time, so exposure of the s a m p l e s to light or bright sunlight should be minimized. There m a y be d i m m i n g of the fluorescence signal of the FLV for incubations exceeding 16 h. It m a y be useful to calculate rates of virus p r o d u c t i o n and r e m o v a l from both the first few time points, and from over the entire course of the experiment. Previous experiments have d e m o n s t r a t e d that in s o m e environments, there a p p e a r s to be an initial 'fast' loss of FLV, followed b y a slower rate of loss over the r e m a i n d e r of the e x p e r i m e n t (Noble and F u h r m a n , 2000). In this case, it m a y be useful to calculate both sets of virus p r o d u c t i o n and r e m o v a l rates to p r o v i d e a range of values.
78
Applications This m e t h o d is unique in that it permits simultaneous determination of rates of virus production and removal using epifluorescence microscopy. For example, an experiment in Southern California coastal waters d e m o n strated rates of viral production and removal of 3.9% h l and 4.5% 11 ', respectively (Noble and Fuhrman, 2000). Experiments performed t h r o u g h o u t the southern California Bight have revealed virus production rates ranging from 2 × 1if' to 3 x 1()1~' virus l ' d ', and indicate that the turnover of virus populations in both nearshore and offshore waters is 1-2 days (Noble and Fuhrman, 2000). Using a range of values from 20 to 50 for burst size, it was estimated that from 24-125% of the bacterial population was killed by viruses in southern California waters (Noble and Fuhrman, 2000). A limitation of the FLV m e t h o d when making measurements in surface waters is the requirement for dark incubations. In Noble and Fuhrman (2000), experiments were started at dusk, so as to provide measurements of in situ rates of virus production and removal. However, processes that affect rates of production and removal in surface waters may be quite different d u r i n g daylight hours. Rates of virus removal could be underestimated, as sunlight is k n o w n to be responsible for a significant portion of degradation and removal of virus particles in seawater (Noble and Fuhrman, 1997). However, simultaneously, sunlight m a y actually be responsible for increased viral production due to prophage induction of lysogenic bacteria and subsequent bacterial cell lysis. The production values could also be overestimated because of diminished bacterial activity in seawater due to intense sunlight (Aas et al., 1996). It should be noted, as discussed in Chapter 3: Enumeration of viruses, that SYBR Gold has been identified as a useful substitute for SYBR Green I as a virus stain (Chen et al., in press). SYBR Gold stained bacteria and viruses are brighter than those stained with SYBR Green I, and SYBR Gold is m a r k e d l y less expensive. Although the authors have not performed a quantitative comparison between the use of SYBR Green I and SYBR Gold for the FLV tracer method, we have every reason to believe that SYBR Gold w o u l d be a viable substitute.
CONCLUSIONS The principles u p o n which each of the methods detailed within this chapter are based are completely different. Therefore, there are advantages and disadvantages unique to each and these have been presented in the corresponding sections. The choice of which m e t h o d to use will d e p e n d upon such factors as experimental design, available equipment, and capability to use radioisotopes. Another consideration is whether it is necessary to estimate the production rate of tile entire viral assemblage or only of bacterial viruses. The FLV tracer dilution m e t h o d targets the entire viral assemblage, whereas the FVIC-based m e t h o d is specific to estimating bacteriophage productivity. Tile radiolabeling method is specific 79
to bacteriophage productivity w h e n using ~H-TdR, but has been used with 32po43 to estimate total viral productivity. The FLV tracer m e t h o d is likely to be more sensitive than the radiolabeling method, which would prove useful in oligotrophic and deep waters. The lowest significant rates measured by radiolabeling are on the order of 10 ~viruses 1 ' d ~in surface waters. Rates at least as low as 1 x lff' viruses 1 ' d ~have been reliably obtained by tracer dilution from waters as deep as 60m. Waters with lower rates of production have not been assayed so the practical lower limit of detection with this m e t h o d is not yet known. However, assuming that FLV abundance can be determined at each time point with _< 20% error, then rates on the order of 10~ viruses 1' d ', or lower, are probably measurable. The sensitivity of the FVICbased m e t h o d is difficult to compare with the others because it depends heavily on the assumptions used for calculation. However, we can make a rough estimate by assuming that (1) only one of 1000 cells examined is infected (FVIC = 0.001), (2) bacterial productivity is reasonably low at 1 x 10~ cells 1' d ~, and (3) burst size is between 10 and 100. Using these assumptions, the m i n i m u m estimated productivity is between 107 and 10 ~ bacteriophage 11 d '. While this m e t h o d is potentially quite sensitive, it is also based on a n u m b e r of u n p r o v e n assumptions. Despite the fundamental differences among these methods, both in terms of underlying assumptions and potential limitations, the results obtained with each have been comparable, within an order of magnitude. Rates determined by the FLV tracer m e t h o d in Southern California coastal waters (Fuhrman and Noble, 2000) were within the range of those obtained by radiolabeling in similar environments (Steward et al., 1992b). In another study, production rates were estimated by FVIC-based and radiolabeling methods on the same samples (Steward et al., 1996). In the five samples assayed, radiolabeling estimates were within the estimated ranges derived from FVIC. In part this was due to the large uncertainty in the FVIC-based estimates, but in the worst case the means differed by less than a factor of four. Measuring viral productivity in complex communities remains a significant challenge. Agreement between any two methods is still marginal and no single method has yet appeared as the gold standard. Nevertheless, estimates obtained by the wide variety of available methods has led to the general conclusion that viruses contribute significantly to microbial mortality and to the cycling of carbon and nutrients in aquatic food webs (Wilhelm and Suttle, 1999).
References Aas, P., Lyons, M., Pledger, R., Mitchell, D. L. and Jeffrey, W. H. (1996). Inhibition of bacterial activities by solar radiation in nearshore waters and the Gulf of Mexico. Aquatic Microbial Ecol. 11, 229-238. Bergh, O., Bgrsheim, K. Y., Bratbak, G. and Heldal, M. (1989). High abundance of viruses found in aquatic environments. Nature 340, 467-468. Binder, B. (1999). Reconsidering the relationship between virally induced bacterial mortality and frequency of infected cells. Aquatic Microbial Ecol. 18, 207-215. Blackburn, T. H. (1979). Method for measuring rates of NH4 turnover in anoxic
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marine sediments, using a '~N-NHa dilution technique. Appl. E11viron. Microbiol. 37, 760-765. B~rsheim, K. Y., Bratbak G. and Heldal M. (1990). Enumeration and biomass estimation of planktonic bacteria and viruses by transmission electron microscopy. Appl. Environ. MicrobioI. 56, 352-356. Bratbak, G., Heldal, M., Norland, S. and Thingstad, T. F. (1990). Viruses as partners in spring bloom microbial trophodynamics. Appl. Environ. Microbiol. 56, 1400-1405. Bratbak, G., Thingstad E and Heldal, M. (1994). Viruses and the microbial loop. Microbial Ecol. 28, 209-221. Bratbak, G., Heldal, M., Thingstad, T. F. and Tuomi, P. (1996). Dynamics of virus abundance in coastal seawater. FEMS Microbiol. Ecol. 19, 263-269. Chen, E, Lu, J. R., Binder, B. J., Liu, Y. C. and Hodson, R. E. Enumeration of marine viruses stained with SYBR Gold: Application of digital image analysis and flow cytometry. Appl. Environ. Microbiol. in press. Fuhrman, J. A. (1987). Close coupling between release and uptake of dissolved free amino acids in seawater studied by an isotope dilution approach. Mar. Ecol. Pros,. Ser. 37, 45-52. Fuhrman, J. A. (1999). Marine viruses and their biogeochemical and ecological effects. Nature 399, 541-548. Fuhrman, J. A. and Azam, E (1982). Thymidine incorporation as a measure of heterotrophic bacterioplankton production in marine surface waters: evaluation and field results. Mar. Biol. 66, 109-120. Fuhrman, J. A. and Noble, R. T. (1995). Viruses and protists cause similar bacterial mortality in coastal seawater. Limnol. Oceanog. 40, 1236-1242. Glibert, E M. (1982). Regional studies of daily, seasonal, and size fractionation variability in a m m o n i u m regeneration. Mnr. Biol. 70, 209-222. Guixa-Boixareu, N., Calder6n-Paz, J. I., HeldaI, M., Bratbak G. and Pedr6s-Ali6, C. (1996). Viral lysis and bacterivory as prokaryotic loss factors along a salinity gradient. Aquatic Microbial Ecol. 11, 215-227. Hara, S., Terauchi, K. and Koike, I. (1991). Abundance of viruses in marine waters: Assessment by epifluorescence and transmission electron microscopy. Appl. Environ. Microbiol. 57, 2731-2734. Heldal, M. and Bratbak, G. (1991). Production and decay of viruses m aquatic environments. Mar. Ecol. Pro% Set'. 72, 205-212. Hennes, K. P. and Simon, M. (1995). Significance of bacteriophages for controlling bacterioplankton growth in a mesotrophic lake. Appl. Environ. Microbiol. 61, 333-340. Hermes, K. P. and Suttle, C. A. (1995). Direct counts of viruses in natural waters and laboratory cultures by epifluorescence microscopy. Limnol. Oceano,¢. 40, 1050 1055. Kepner, R. L. J., Wharton, R. A. and Suttle, C. A. (1998). Viruses in Antarctic lakes. Limlml. Oceanog. 43, 1754 1761. Moriarty, D. J. W. (1986). Measurements of bacterial growth rates in aquatic systems from rates of nucleic acid synthesis. In: AdvamTes in Microbial Ecolosy (K.C. Marshall, Ed.), pp. 245-292. Plenum Press, New York. Noble, R. T. and Fuhrman, J. A. (1997). Virus decay and its causes in coastal waters. Appl. Environ. Microbiol. 63, 77-83. Noble, R. T. and Fuhrman, J. A. (1998). Use of SYBR Green 1 for rapid epifluorescence counts of marine viruses and bacteria. Aquatic Microbial Ecol. 14, 113 118. Noble, R. T. and Fuhrman, J. A. (2000). Rapid viral production and removal as measured with fluorescently labeled viruses (FLV) as tracers. Appl. Environ. Microbiol. 66(9), 3790-3797.
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Proctor, L. M. and Fuhrman, J. A. (1990). Viral mortality of marine bacteria and cyanobacteria. Nature 343, 60-62. Proctor, L. M., Okubo, A. and Fuhrman, J. A. (1993). Calibrating estimates of phage induced mortality in marine bacteria: Ultrastructural studies of marine bacteriophage development from one-step growth experiments. Microbial Ecol. 25, 161-182. Steward, G. E and Azam, E (2000). Analysis of marine viral assemblages. In: Microbial Biosystems: New Frontiers, 8th International Symposium for Microbial Ecology (C. R. Bell, M. Brylinsky and P. Johnson-Green, Eds), Atlantic Canada Society for Microbial Ecology, Halifax, Canada, 159-165. Steward, G. E, Wikner, J., Smith, D. C., Cochlan, W. P. and Azam, E (1992a). Estimation of virus production in the sea: I. Method development. Mar. Microbial Food Webs 6, 57-78. Steward, G. E, Wikner, J., Cochlan, W. P., Smith, D. C. and Azam, E (1992b). Estimation of virus production in the sea: ll. Field results. Mar. Microbial Food Webs 6, 79-90. Steward, G. E, Smith, D. C. and Azam, E (1996). Abundance and production of bacteria and viruses in the Bering and Chukchi Sea. Mar. Ecol. Prog. Set. 131, 287-300. Steward, G. E, Elola, J. and Azam, E (2000). Genome size distributions indicate variability and similarities among marine viral assemblages. Limnol. Oceanog. 45, 1697-1706. Weinbauer, M. G. and H6fle, M. G. (1998). Significance of viral lysis and flagellate grazing as factors controlling bacterioplankton production in a eutrophic lake. Appl. Environ. Microbiol. 64, 431-438. Weinbauer, M. G. and Peduzzi, P. (1994). Frequency, size and distribution of bacteriophages in different marine bacterial morphotypes. Mar. Ecol. Pro,g. Set. 108, 11-20. Weinbauer, M. G. and Suttle, C. A. (1997). Comparison of epifluorescence and transmission electron microscopy for counting viruses in natural marine waters. Aquatic Microbial Ecol. 13, 225-232. Weinbauer, M. G., Fuks, D. and Peduzzi, P. (1993). Distribution of viruses and dissolved DNA along a coastal trophic gradient in the Northern Adriatic Sea. Appl. Environ. Microbiol. 59, 4074-4082. Wells, M. L. and Goldberg, E. D. (1992). Marine submicron particles. Mar. Chem. 40, 5-18. Wikner, J., Vallino, J. J., Steward, G. E, Smith, D. C. and Azam, E (1993). Nucleic acids from the host bacterium as a major source of nucleotides for three marine bacteriophages. FEMS Microbiol. Ecol. 12, 237-248. Wilhelm, S. W. and Suttle C. A. (1999). Viruses and nutrient cycles in the sea. Bioscience 49, 781-788. Xenopolous, M. A. and Bird, D. E (1997). Microwave enhanced staining for counting viruses by epifluorescence microscopy. Limnol. Oceanog. 42, 1648-1650. Zar, J. (1996). Biostatistical Analysis, third ed., Prentice Hall.
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List of suppliers Advantec MFS, Inc. 6691 Owens Drive Pleasanton, CA 94588-3335, USA Tel: +1 800 334 7132 or +1 925 225 0349 Fax: +1 925 225 0353 E-maih
[email protected]
Stainless steel filter holders, stainless steel pressure filtration vessels A/G Technology Corporation 101 Hampton Avenue Needham, M A 02494-2628, USA Teh +1 781 449 5774 or +1 800 248 2535 Fax: +1 781 449 5786 E-mail:
[email protected] lnternet: www.a~tech.com
Millipore, Inc. 80 Ashby Road Bedford, M A 01730-2271, USA Teh +1 781 533 6000 Fax: +1 781 533 3110 lnternet: www.millipore.com
Ultrafiltration products (Millipore and Amicon brands), membrane filters Molecular Probes, Inc. 4809 Pitchford Avenue Eugene, OR 97405-0469, USA Teh +1 541 465 8300 Fax: +1 541 344 6504 E-maih
[email protected] Internet: www.probes.com
SYBR Green I and SYBR Gold stains
Ultrafiltration products Amersham Pharmacia Biotech AB SE-751 84 Uppsala, Sweden TH: +46 (0) 18 612 O0 O0 Fax: +46 (0) 18 612 12 O0 lnternet: www.apbiotech.com
Radiochemicals ([~H]-TdR)
NEN "~'Life Science Products, Inc. 549 Albany Street Boston, M A 02118-2512, USA Teh +1 617482 9595 or +1 800 551 2121 Fax: +1 617482 1380 lnternet: "~Lv~U'~U.tl~'II/ffl~SCJ.COtll
Radiochemicals ([~H]-TdR) Fisher Scientific, Inc. 2000 Park Lane Pittsbm2~h, PA 15275, USA Teh +1 800 766 7000 Fax: +1 800 926 1166 Internet: w w w l .fishersci.com
General laboratory supplies and equipment, plasticware, chemicals, Acrodisc filters (Gelman)
Sigma-Aldrich, Inc. 3050 Spruce Street St Louis, M O 63103, USA Tel: +1 314 771 5765 o7" +1 800 325 8070 Fax: +1 314 771 5757 E-maih
[email protected] Internet: www.sigma-aldrich.com
Enzymes, chemicals, biochemicals, p-phenylenediamine
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Ted Pella, Inc. P.O. Box 492477, Redding, CA 96049-2477, USA Teh +1 530 243 2200 or +1 800 237 3526 Fax: +1 530 243 3761 E-maih
[email protected] Internet: www.tedpella.com
Electron microscopy supplies
Whatman International Ltd Catalogue Sales Department St Leonard's Road 20/20 Maidstone Kent ME16 OLS Teh +44(0)1622 674821 Fax: +44(0)1622 682288 E-mail:
[email protected] lnternet: www.whatman.com
Direct purchase of Anodisc filters VWR Scientific Products, Inc. V W R International 3000 Hadley Road So. Plainfield, NJ 07080, USA Teh +1 800 932 5000 or +1 908 757 4045 Fax: +1 908 757 0313 lnternet: www.tdwrs]d.cotn
Anodisc filter membranes, slides, cover slips
Worthington Biochemical Corporation 730 Vassar Avenue Lakewood, NJ 08701, USA Tel: +1 732 942 1660 or +1 800 445 9603 Fax: +1 732 942 9270 or +1 800 368 3108 E-mail:
[email protected] lnternet: www.worthingtonbiochem.com
Enzymes
84
6 Fingerprinting Viral Assemblages by Pulsed Field Gel Electrophoresis (PFGE) Grieg F Steward Monterey BayAquarium Research Institute,Moss Landing,California, USA
CONTENTS Introduction
Principle Application Conclusion
ee4,eee
INTRODUCTION Viruses are the most abundant microorganisms in marine and freshwater environments and perhaps the most genetically diverse (Fuhrman and Suttle, 1993). Counting viruses in aquatic samples is now a routine matter, but assessing the diversity and dynamics within complex assemblages is still a challenge. DNA-based fingerprinting approaches which rely on amplification of rRNA gene fragments by PCR have facilitated analyses of bacterial community composition. These approaches have more restricted application when analyzing viral assemblages, because of the extreme genetic diversity among viruses. Unlike bacteria, there are no gene sequences conserved in all viruses which can serve as universal primer sites for PCR amplification. PCR-based analyses of viral assemblages must therefore target specific subsets of the total viral assemblage. For example, PCR amplification of specific genes has recently been used to examine the genetic diversity among cyanophages (Fuller et al., 1998) and among phycodnaviridae (Chen et al., 1996; Short and Suttle, 1999). A more general fingerprinting approach, which encompasses the total viral assemblage, can be a valuable complement to these more specific, higher resolution analyses. The approach described here uses variation in genome size as the basis for obtaining a fingerprint of a viral assemblage (Klieve and Swain, 1993). A whole genome fingerprinting approach is possible, because viral genomes can vary greatly in length (a few thousand to hundreds of thousands of base pairs) yet they fall within a range that is easily resolved using pulsed field gel electrophoresis (PFGE). The
METI IODS IN MICROBIOLOGY, VOLUME 30 ISBN 0-12-521530-4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any form reserved
PFGE fingerprinting technique provides a quick and relatively simple means of visualizing differences in the composition of viral assemblages (Swain et al., 1996; Wommack et al., 1999a; Steward et al., 2000). As a supplement to the more specific treatment of PFGE provided in this chapter, the reader is encouraged to consult the excellent introductory text to PFGE by Birren and Lai (1993).
e,e, e e e , e, P R I N C I P L E Viruses are harvested from a water sample by ultrafiltration and the capsids are destabilized to release the viral DNA. Intact viral genomes are separated by size in an agarose gel by PFGE. After separation, the DNA banding pattern is revealed with a fluorescent DNA stain. The banding pattern provides a visual record of the genome size distribution which can be used for qualitative and quantitative comparisons among samples. Using image analysis, the molecular weight and mass of DNA in each band are determined by comparison with the migration rate and fluorescence intensity of DNA standards. Molecular weight and mass are then used to calculate the genome copy number in each discrete band or molecular weight size range.
Equipment and reagents
•
•
Tangential flow ultrafiltration system: A tangential flow system is necessary if viruses are to be harvested from a liter or more of water. Choose a system with the smallest minimum recirculation volume which will still provide a reasonable processing time (i.e. less than an hour or two). A system with smaller tubing and a smaller membrane surface area will process more slowly, but will also have a smaller minimum recirculation volume. This means the concentration factor achievable for a given initial sample volume will be greater. Spiral wound membranes - - designed for processing low viscosity, particle-free fluids - - are well suited to harvesting viruses from prefiltered (0.2 lum) water. Other configurations, however, such as membrane cassettes or hollow fiber cartridges, and other systems such as vortex flow filtration, can also be used. A 30 000 molecular weight cutoff (MWCO) is recommended to ensure retention of smaller viruses, but 100000 MWCO membranes may be adequate. Membranes rated at < 30 000 MWCO will filter more slowly and retain more unwanted low molecular weight material. Sterile flters (0.2tim pore size): Sterivex GV filters (Millipore, Bedford, Massachusetts, USA) or their equivalent are adequate for filtering 10 ml to I 01 of water depending on the particle load. Filters with larger surface area (e.g. pleated capsule filters) are required for larger volumes (ten to hundreds of liters). *Centrifugal ultrafiltration units (100 000 MWCO): Large capacity (20 to 80 ml) units are useful as a secondary concentration step.They can also be used for primary concentration of small volume samples (< I liter). Small capacity (0.5 ml) centrifugal ultrafiltration units are used for final virus concentration and preparation of viral DNA. 86
•
• •
•
• • • • • •
•
• • • •
*Centrifuge: Moderate speed (capable of 4000g) with swinging buckets and adapters to hold large capacity centrifugal ultrafiltration units. A centrifuge with refrigeration is recommended. *Microcentrifuge: Speed must be adjustable. A unit with refrigeration is recommended. Pulsed field gel electrophoresis system: The system should be capable of providing resolution of D N A up to several hundred-thousand base pairs.The procedures described in this chapter are for units using a clamped homogeneous electric field (CHEF) configuration which are available commercially from Bio-Rad. Gel documentation equipment:A variety of system configurations are possible. Laser gel scanners are preferred as they provide the greatest sensitivity and resolution and direct acquisition of digital images. For systems using illumination by UV lamps, epi-illumination is reported to provide higher sensitivity than transillumination for gels stained with SYBR Green I (Molecular Probes, Inc.). Regardless of illumination method, documentation with a digital camera is preferred over film since the images can be directly imported into gel analysis programs. If film must be used, a scanner can be used to convert the photographs into digital images, but the dynamic range and resolution will be lower than for the direct, digital image acquisition. Gel analysis software:The software should have capabilities for band recognition, calculation of integrated intensity, and molecular weight determination. Fluorescent DNA stain for D N A quantification such as PicoGreen (Molecular Probes). Fluorometer capable of measuring fluorescence of stained D N A (e.g. 502 nm excitation, 523 nm emission peaks for PicoGreen). Purified Bacteriophage lambda DNA. Pipets, micropipettors. Agarose: A PFGE-grade agarose with a standard gelling temperature such as SeaKem* Gold (FMC Bioproducts) can be used for routine application. If D N A is to be recovered from the gel following electrophoresis, then a low-meltingpoint agarose such as SeaPlaque GTG ~' (BioWhittaker) should be used instead. A low-melting-point agarose is also recommended for embedding samples in agarose plugs prior to electrophoresis. DNA molecular weight standards: Standards should cover the range from about ten- to several hundred-thousand base pairs. Low and midrange PFG Markers (New England BioLabs) are convenient as they each provide full range coverage with a single marker. Other useful markers are a 5 kb ladder (concatemers of a 4.8kb plasmid) and a lambda ladder (concatemers of lambda phage genomes) which can be used in combination (available from various suppliers). DNA mass standards: D N A mass ladder (GIBCO/BRL) or dilutions of lambda phage DNA. Running buffer ( I 0×TBE stock contains per liter: 108 gTris base, 55 g boric acid and 40 ml of 0.5 M EDTA, pH 8). Loading buffer (10× stock contains 25% ficoll, 0.25% Bromophenol blue or xylene cyanol). SYBR Green I (10 000× stock; Molecular Probes) or ethidium bromide (5 mg ml ' stock).
87
iii
. m
• •
Sodium azide: 10% stock solution in water, filtered (0.2 lum). SM or Marine SM (MSM): These are storage buffers for non-marine or marine
bacteriophages. SM contains 100 mM NaCI, 10 mM MgSO4,50 mMTris (pH 7.5), and 0.01% gelatin (Sambrook et al., 1989). MSM is a modification of the original SM recipe which more closely matches the ionic composition and pH of seawater and contains 450 mM NaCI, 50 mM MgSO4, 50 mM Tris (pH 8.0), and 0.01% gelatin. Sterilize by autoclaving and store at room temperature.These buffers can also be prepared without the gelatin for situations where adding additional, high molecular weight protein to the sample is undesirable.
*Alternative: The final virus concentration step m a y be accomplished by ultracentrifugation instead of centrifugal ultrafiltration. In this case an ultracentrifuge and appropriate tubes are required and could replace the centrifuge, microcentrifuge and centrifugal ultrafiltration devices.
Assay In addition to the following detailed description, an overview of the viral fingerprinting procedure is presented as a flow chart (Figure 6.1). Virus concentration and storage
The sample volume required can vary greatly depending on the initial concentration of viruses, losses during processing, sensitivity of detection, and whether extra DNA is desired for multiple gel runs, archiving, or other analyses. For a typical seawater sample, at least 10" viruses or 50 ng of viral DNA are needed to obtain a single fingerprint in a 10 m m wide well. For a typical surface seawater concentration of 10 '~ viruses 1 ', this translates into a m i n i m u m sample volume for a single fingerprint of roughly 100 ml. A larger volume is recommend, however, to account for losses and to have extra material. In practice, process volumes m a y range from around 10 ml to > 50 1, with 1-3 liters being sufficient in most cases.
Processing large volumes (> I liter) Filter sample (0.2 lum pore size) to remove bacteria. For water with a high particle load, prefiltration through a filter with a larger pore size may be useful to avoid clogging the 0.2 lum filter. Concentrate viruses in the filtrate by tangential flow ultrafiltration to < 400 ml then proceed with small volume concentration (below).
Processing small volume samples or primary concentrates (< 400 ml) Q
For primary viral concentrates skip to the next step. For small samples, filter through a 0.2 micron Sterivex filter (Millipore) via syringe or peristaltic pump to remove bacteria. Concentrate viruses by centrifugal ultrafiltration to _<2501ul. Samples exceeding the capacity of the filter reservoir are concentrated by repeated rounds of centrifugation. After each round, the filtrate is discarded and the upper reservoir is refilled with additional sample.
88
Remove Bacteria • Filtration (0.2 pm)
Storage Concentrate Viruses
• Ultrafiltration
• MSM or SM • Na azide
- - ~
• <_4oC
Release DNA
• Embed viruses in agarose • Treat with TE + SDS + Proteinase K OR overnight, room temperature
• Exchange medium with TE (centrifugal ultrafiltration) • Heat to 60°C, 10 min
I I
Storage • TE • 4oc
(longer term)
Separate Genomes by PFGE I _.1~ ~l • 1% agarose ~_ • 0.5x TBE, 14°C • 6 Vcm -~,18 h • Pulse ramp 1 to 10 s
[
Storage • TE • 4oc
(short term only)
Stain and Image Gel
• SYBR Green I • laser gel scanner
Analyze Images
• Identify and quantify bands • Calculate band MW and genome copy number
Descriptive and Comparative Statistics
• Genome size distribution (1 sample) • Similarity coefficients (2 samples) • Cluster analysis (_>3 samples)
Figure 6.1. Flow chart illustrating the major steps in generating and analyzing whole-genome fingerprints of viral assemblages. Bulleted details within each box are those recommended and presented in the text, but alternatives and variations are possible at each step. Dashed arrows represent alternative pathways for temporary storage of samples.
•
•
Recover sample by inverting the filter cup into a collection tube and centrifuging at 250g for I min. Rinse membrane with 250 I~1 of MSM, 0.2 Ium filtered sample, or ultraflltrate, recover and pool. Add sodium azide to 0. 1% final concentration and store at 4°C in the dark.
89
Alternatively, viruses can be concentrated from small volumes by ultracentrifugation. The centrifugation time and speed required will vary d e p e n d i n g on the rotor. Run conditions are calculated from the k factor of the rotor and an assumed sedimentation coefficient of about 100S in order to pellet the smallest viruses. The formula is t = k/S, where t is time in hours, k is the k factor and S is the sedimentation coefficient in Svedbergs. The k factor assumes sedimentation of particles in pure water at 25°C. The time may need to be corrected for salinity and temperature effects on the density and viscosity of water. For example, if viruses are pelleted from seawater not freshwater, centrifugation time should be increased by about 12c7~. Temperature effects on water viscosity become pronounced below 20°C, so centrifugation time should also be increased by an additional 25% for each 5°C decrease below 20°C (Suttle, 1993).
Preparation of viral D N A for electrophoresis There are two approaches to DNA preparation; in agarose plugs or in solution. Traditionally, samples for PFGE are e m b e d d e d in agarose plugs and treated with a chelator (EDTA), detergent, and proteinase K (Klieve and Swain, 1993; Wommack et al., 1999a). However, if the samples are handled carefully, e m b e d d i n g is not necessary for DNA in the size range of typical viral genomes. In addition, proteinase K digestion is not strictly necessary since viral capsids can be sufficiently destabilized and the D N A released by exposure to EDTA and moderate heating. DNA in solution 1.
2.
3. 4.
5. 6. 7.
8.
Transfer an aliquot of viral concentrate containing 40-100 ng of DNA (1 to 2 x 10" viruses) to a Microcon 100 and concentrate to near dryness at 1000g. Required centrifugation time is typically 5 to 20 rain. Rinse the Microcon by gently adding 50 lul of l x TE then concentrating to near dryness (ca. 5 lJl) as above. Centrifugation time is usually 5 to 10 min. Repeat rinse twice more. A d d 10 t~l TE to the Microcon (be sure it wets the membrane) and recover by inverting the filter cup in a collection tube and centrifuging at ca. 250g for 1 min. Repeat the recovery step once or twice more pooling all material into the same collection tube. Heat the recovered sample to 60°C for 10 rain. Cool the sample on ice and spin d o w n any condensation in the tube by brief centrifugation. (Optional). For precise control on the a m o u n t of DNA loaded per lane, determine the DNA concentration in the viral concentrate using the PicoGreen DNA quantification assay (Molecular Probes) and a fluorometer. For the assay, dilute 1-5 1-ll of the recovered DNA into l x TE and proceed according to the manufacturers protocol. Dilute samples as necessary with an appropriate v o l u m e of l x TE then add 1/10 v o l u m e of 10x loading buffer. For m a x i m u m band sharpness, the final sample v o l u m e should be close to the well capacity.
90
9. Gently mix s a m p l e and loading buffer. An effective, gentle mixing m e t h o d is to roll the tube back and forth b e t w e e n t h u m b and forefinger while s i m u l t a n e o u s l y inverting the tube five to ten times. The s a m p l e should stay at the b o t t o m of the tube. Do not vortex. D N A in agarose plugs
1. Prepare a viral concentrate of about 1-5 x 10 Hviruses ml ' (5-25 p g viral D N A ml '). Concentrates m a y be p r e p a r e d by centrifugal ultrafiltration or by ultracentrifugation. Recover concentrated viruses in a small v o l u m e of SM or MSM (without gelatin). If using ultracentrifugation, add the buffer to the viral pellet, cover the tube with Parafilm, and let it sit several hours overnight at 4°C. Pipet in and out to resuspend then transfer to a microcentrifuge tube. If using centrifugal ultrafiltration, concentrate and recover as described in Step 4 of the previous section, substituting SM or MSM (without gelatin) for the TE. 2. Melt a stock of 1.5% (w:v) low-melting-point agarose in SM or MSM (without gelatin) using a m i c r o w a v e oven or hot w a t e r bath. 3. Transfer aliquots of agarose to microtubes in a heating block or water bath at 50 to 60°C to keep them molten. 4. C o m b i n e equal v o l u m e s of viral concentrate (at room temperature) and molten agarose, vortex or pipet in and out several times to mix, then quickly transfer to plug molds and let them solidify. 5. Push the plugs into a microcentrifuge tube containing 3 to 5 v o l u m e s of freshly p r e p a r e d extraction buffer (100 mM EDTA, p H 8.0, 1% SDS and 1 m g ml ' proteinase K). 6. lncubate overnight at r o o m t e m p e r a t u r e with gentle agitation. 7. Rinse the plugs 3 x 30 min with l x TE. 8. Plugs can be stored at 4°C in l x TE. 9. To prepare a p l u g for electrophoresis, place it on a piece of Parafilm, blot a w a y excess liquid and trim with a razor blade so it will fit within a gel well. D N A size and mass markers
A n u m b e r of size m a r k e r s for pulsed field gel electrophoresis are commercially available. The c o m m o n l y available 5 kb and l a m b d a ladders can be used in combination to cover the range of viral g e n o m e sizes. The l a m b d a ladder is supplied in agarose plugs while the 5 kb ladder is in solution. The l a m b d a ladder is at a sufficiently high concentration that only a small sliver of the p l u g is needed (< 1 mm). This leaves e n o u g h r o o m that the 5 kb ladder can be loaded into the s a m e well. Full size range coverage can then be achieved in a single lane. Specially designed ladders which s p a n the entire range of viral g e n o m e sizes are also available ( N e w England BioLabs). At least two size-marker lanes should be run, one at either end of the gel. For precise molecular weight determinations, a m a r k e r lane in the m i d d l e of the gel m a y be needed in s o m e cases. To quantify D N A based on fluorescence intensity, D N A m a s s standards should also be run to compensate for any nonlinearity in fluorescent signal detection. Mass ladders
91
are commercially available which allow calibration to be carried out using only a single gel lane. However, these mass ladders are usually comprised of small DNA fragments (e.g. 1 to 10 kb for the High DNA Mass TM Ladder; Life Technologies Inc.). Most of the small fragments will run off the gel using the PFGE conditions typical for viral fingerprinting. This can be prevented by loading the mass ladder in the middle or near the end of the run. Alternatively, the mass ladder or a series of dilutions of lambda DNA could be run separately by conventional electrophoresis in a gel of the same thickness and percent agarose. The gels should then be stained simultaneously in the same batch of stain and p h o t o g r a p h e d together.
Running the gel 1. Cast a 1% agarose gel in 0.5x TBE. A comb with 5 to 10-ram-wide wells
2.
3. 4.
5.
6.
is recommended. Wider wells make it easier to distinguish faint bands from background. First load any samples which are e m b e d d e d in plugs. Before inserting a plug, fill the well with running buffer to ensure no air bubbles become trapped in the well. For the lambda ladder marker, cut off a thin sliver with a razor blade then place the sliver (about 0.5 m m x 0.5 m m x 5-10 mm) against the side of the well. Small spatulas or plastic inoculating loops are convenient for handling gel plugs and slices. Place the gel in the electrophoresis chamber, add running buffer slowly to avoid dislodging the gel from the casting plate, and start recirculating and chilling the buffer. The chiller should be adjusted so that the buffer temperature in the chamber is 14°C. Recirculate the buffer at a rate of about 0.5 to 1 1 min ' (ca. 1 tank v o l u m e every 2 to 3 minutes). Note: To save time, the chamber can be pre-filled and the buffer chilled while the gel is solidifying. When lowering the gel into the buffer, care must be taken not to dislodge the gel from casting plate. Load liquid samples and markers, pipetting slowly to avoid shearing the DNA. Run the gel at 6 V cm ~for 18 h with pulses ramping from 1 to 10 s at a reorientation angle of 120 °. These conditions provide good separation from 10 to ca. 200 kb. Pulse and run time m a y be varied to improve separation in specific size ranges. Others have used a pulse ramp of 1 to 15 s over a run time of 22 h (Wommack et al., 1999a). Longer pulse and run times improves separation of higher molecular weight material. If pulse and run times are increased too much, however, resolution in the lower molecular weight regions will be sacrificed. Consult Birren and Lai (1993) for a more detailed discussion of the effects of switch and run times on DNA separations. Stain the gel in the dark for 0.5 to i h in about 150 to 200 ml of running buffer containing SYBR Green I (lx) or ethidium bromide (0.05 tJg ml '). If using ethidium bromide, then destain the gel for about 0.5 h in 5 to 10 gel-volumes of water. Acquire an image of the gel with a gel documentation system. If using film, several exposures m a y be useful to optimize the n u m b e r of bands 9:l
within the dynamic range of the film. Laser scanners should be set to high resolution to improve band discrimination.
Analysis of banding patterns Description of an individual sample Quantitative analyses of banding patterns are possible with a variety of commercially available software programs. The program should have the capability to (1) identify bands (with adjustable detection parameters and manual editing capability), (2) integrate the fluorescence intensity of individual and partly overlapping bands, and (3) calculate molecular weight and mass for each band relative to migration distance and intensity of standards on tile same gel. This information along with some simple spreadsheet calculations will provide a statistical description of a given sample in terms of the range, mean, median, and mode of the detectable viral genome sizes. The data can also be graphically displayed as a size-frequency distribution. The calculations for determining the fraction of virus-like genomes within a given band or within size-range bins are presented below. Note that DNA size standards are usually reported in terms of length (base pairs) rather than molecular weight, which is used in the equations here. DNA size in base pairs must first be converted to molecular weight (M) using: M = S x MI, P
(6.1)
where S is the size in base pairs and M,., is average molecular weight of a base pair ( = 660). Genome copy number in each of the detected bands is then calculated as: Q
Di = -x Av
(6.2)
Mi where C, is the genome copy number, D, is the mass (in grams) and M, is molecular weight (in Da) of the DNA in band i, and Av is Avogadro's number (6.022 x 10>). The absolute number of genomes is directly dependent on the amount of sample loaded on the gel. Therefore, to facilitate comparisons among samples, copy number is better expressed as a relative abundance. The fraction of the total viral genomes present in each of the bands is calculated as Ci I
-
~7
Y_.Ci ;= 1
93
(6.3)
where F, is the fraction of genomes and C, the genome copy number in band i, and n is the total number of bands. These data can be graphically presented as size-frequency plots (Steward and Azam, 2000). In m a n y cases, there are regions on the gel where individual bands cannot be resolved. To deal with this, as well as to standardize comparison among different samples, it m a y be preferable in some instances to bin the frequency data into defined size ranges and plot as a histogram (Steward et al., 2000). The average genome size for an assemblage can be derived from these data by first calculating an abundance-weighted molecular weight (M*) for each band i as M7 = M i x C i
(6.4)
The average genome size (G.~v~)can then be calculated as
(6.5)
G a v g _ ,=1 i-1
q
The mode of the genome size distribution is readily identified as the genome size or (size bin) with the m a x i m u m F~. The median genome size can also be identified by calculating a running sum of F, from the largest to the smallest genome (or vice versa). The genome size or size class at which the sum is at, or closest to, 50% is the median.
Comparisons among samples A similarity coefficient can be calculated for any two samples from their genome size-frequency distributions. In the case of viral fingerprints, frequency data are expressed as proportions rather than absolute abundance (from Equation (6.3)). Two measures of similarity designed to handle proportional data are the Renkonen index (percentage similarity) and the simplified Morisita index (Krebs, 1999). Binary similarity coefficients can also be calculated by considering only the presence or absence of different-sized genomes (Wommack et al., 1999a). Since information on relative abundance is ignored, this provides a much cruder estimate of similarity. However, it m a y be preferred in situations where the relative recoveries of different viruses is suspected or known to vary significantly from sample to sample. In this case, changes in the relative brightness of different bands could be due to artefacts of sample processing and are best not considered. Relationships among three or more samples can be analyzed by cluster analysis. The most frequently used strategy for clustering is the unweighted pair-group method using arithmetic averages (UPGMA). The 94
algorithm makes use of a matrix of similarity coefficients obtained from pairwise comparisons of all samples. This is a hierarchical, agglomerative approach that is used to create dendrograms illustrating the relative similarities among groups of samples. The details of the calculation of similarity coefficients and of cluster analysis are available from many sources and are not reproduced here. For one recent, lucid presentation and discussion of these methods the reader is referred to Krebs (1999).
Troubleshooting Migration is too fast or too slow
The recommended electrophoresis parameters apply for ca. 21 of 0.5x TBE in a CHEF unit (Bio-Rad) run at 14°C. If other buffer formulations, volume or temperatures are used, the parameters may need adjusting. For example, DNA will migrate faster in Tris-acetate EDTA (TAE) buffer than in TBE, but TBE is often used because it has a greater buffering capacity. Running gels at lower temperatures improves band resolution, but dramatically reduces the migration rate. Migration rate will also decrease with increasing buffer volume and increasing ionic strength. For a comprehensive discussion of these and other factors influencing the migration rate of DNA in pulsed field gels the reader is referred to Birren and Lai (1993). Migration is uneven across the gel
Uneven migration is most likely the result of an improperly leveled gel box. The bottoms of some electrophoresis tanks can have some curvature making small, bubble levels less accurate. For best results use a bubble level which is short enough to fit in the tank, but long enough to rest on electrodes at either side of the tank. Alternatively, the tank level can be checked by adding a small amount of buffer to the chamber then tipping the chamber up, first on the front two feet then the back two feet. With the chamber tipped, adjust the feet so that the buffer level is at the same position relative to the electrodes on opposite sides of the tank. Degraded D N A
Degraded DNA may be caused by nucleases or mechanical shearing and will appear as a smear on the gel. DNA released from viral capsids by EDTA and heat treatment tends to degrade with time in solution. Therefore, samples to be loaded in solution should be prepared fresh from the viral concentrates and should be mixed or pipetted gently to prevent mechanical shearing. If one wishes to store samples as DNA rather than as viral concentrates, the viruses should be embedded and the DNA extracted and stored in agarose plugs rather than in solution. Band resolution is somewhat lower for embedded DNA (see Applications), but the DNA will be better preserved. Remember to wear gloves whenever 95
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, m
working with DNA to minimize the potential for DNase contamination. In addition, be sure the DNA storage buffer contains sufficient EDTA to chelate any magnesium ions as this will inhibit DNase. On the other hand, if storing intact viruses, a buffer containing magnesium ions (like SM or MSM) is used in order to maintain stability of the viral capsids. A preservative such as 0.1% sodium azide should be added to prevent the growth of bacterial contaminants. Both DNA and viruses should be stored cold (4°C) and dark. Viral concentrates can also be stored frozen, but there are no reports yet on the relative stability of frozen vs. refrigerated concentrates of natural viral assemblages. Viral counts do decline with time in refrigerated concentrates (unpublished observation) so samples should be analyzed as soon as possible.
Artefactual banding patterns If a viral concentrate is stored improperly, the banding pattern could be compromised. Banding patterns from seawater samples are typically complex. If only one or two bands dominate a sample, it could be the result of an improperly stored concentrate. An example of this is shown in Figure 6.2. Two filtered (0.2 lJm), unpreserved primary concentrates were stored at 4°C for about 3 weeks. Inspection of the samples by epifluorescence microscopy showed bacterial contamination in both. In addition, one of the samples showed a _> 12-fold increase in viral abundance. The PFGE banding pattern of that sample was dominated by one band indicating that one of the bacterial contaminants served as a host for viral replication during storage. Banding patterns could also be compromised by differential degradation of virus strains within a sample during storage. As mentioned above, viruses do decay with storage, but the degree to which this affects the viral fingerprint has not yet been tested. Dominance of one or a few bands is not necessarily an artefact as it could also occur as a result of natural processes, for example, by mass viral lysis of a monospecific bloom of some organism. In order to distinguish real dominance from artefact it is helpful to monitor the recovery of viruses at each step of harvesting and concentration (e.g. by epifluorescence microscopy, Chapter ll). If recovery is unusually low, or impossibly high, or if a growth of bacterial contaminant is detected during storage, the resulting fingerprint should be considered suspect.
APPLICATION Genomic fingerprinting of viral assemblages by pulsed field gel electrophoresis may be applied to essentially any type of sample from which a sufficient number of viruses can be harvested. So far the technique has been used to analyze phage communities in the rumen of sheep (Klieve and Swain, 1993) and viral assemblages in estuarine (Wommack et al., 1999a; 1999b) and oceanic waters (Steward and Azam, 2000; Steward et a]., 2000). Preliminary data show that the technique can also be applied to 96
M
1
2
194 kb 146 kb 96 kb
48 kb L.
;>9 kb i,.e~
Figure 6.2. Example of an artefactual banding pattern caused by storage of an unpreserved sample for three weeks at 4°C. Viruses were concentrated from surface waters of Monterey Bay, CA and separated by PFGE using a 1-6 s pulse ramp over 16 h. Other electrophoresis conditions are as described in the text. Lane 1 shows a typical banding pattern for one sample which had bacterial contamination, but showed no increase in viral abundance during storage. Lane 2 shows a sample which had bacterial contamination and showed > 12-fold increase in viral abundance during storage. Lane M is a marker lane containing a 5 kb ladder and a lambda ladder with the size of selected bands indicated on the left.
viruses harvested from sea ice a n d s e d i m e n t pore waters (unpublished observations). A c o m p a r i s o n of various s a m p l e p r e p a r a t i o n procedures is illustrated using surface seawater from Monterey Bay (Figure 6.3a). Concentration b y centrifugal ultrafiltration or by ultracentrifugation yielded essentially the s a m e results, but the ultracentrifuged s a m p l e s h o w e d some d e g r a d a t i o n (smearing in the lower molecular weight range). This p r o b a b l y reflects limited mechanical shearing d u e to the gentle s a m p l e agitation which was used w h e n releasing D N A from the viral pellet. Banding patterns were also similar w h e t h e r D N A w a s p r e p a r e d in agarose plugs or in solution, although band resolution is s o m e w h a t poorer for e m b e d d e d samples. This difference in resolution between samples loaded in solution or as gel plugs is also illustrated with l a m b d a p h a g e D N A (Figure 6.3a). Note that for e m b e d d e d samples, b a n d sharpness is limited by the thickness of the gel plug. Although concentration of viruses from 3 to 10 1 of w a t e r is r e c o m m e n d e d in order to have plenty of extra material, small-scale concentrations can also p r o v i d e e n o u g h material for
97
(~) M
1
2
3
365 kb 225 kb
(b) M
4
5
M
1
2
194 146
96 86 kb 58 kb
48
29
29 kb
Figure 6.3. Comparison of sample preparation procedures using surface seawater from Monterey Bay, CA and bacteriophage lambda DNA. Pulse conditions were a 1-t0 s ramp over 18 h and sizes of selected marker bands are indicated on tile left. (A) An initial viral concentrate was prepared from seawater by tangential flow ultrafiltration. The viral DNA was then processed for PFGE by one of three methods: Lane 1 - - viruses were further concentrated by centrifugal ultrafiltration, embedded in agarose, treated with EDTA, SDS and proteinase K, then loaded as a gel plug. Lane 2 viruses were concentrated as for Lane 1, but were diluted in TE and heated to release DNA, then loaded as a solution. Lane 3 - - as in Lane 2, except that viruses were concentrated by pelleting in an ultracentrifuge instead of using centrifugal ultrafiltration. Bacteriophage lambda DNA was loaded in a gel plug (lane 4) or in solution (lane 5). Lanes labeled 'M' contain a 5 kb ladder and a Saccharomyces cerevisiae chromosomal DNA marker (BioRad). (B) Viruses were concentrated from 80ml of 0.2urn-filtered seawater (20m depth) using a Centricon 20 (lane 1) or Centricon 80 (lane 2); all of each concentrate was then processed for electrophoresis on Microcon 100 centrifugal units and loaded on the gel as described in the text. Marker Lane M contains a 5 kb ladder and a lambda ladder (BioRad).
a f i n g e r p r i n t in s o m e e n v i r o n m e n t s . F o r e x a m p l e o n l y 80 m l of s e a w a t e r f r o m 20 m d e p t h in M o n t e r e y B a y w a s sufficient to o b t a i n a v i r a l fingerp r i n t ( F i g u r e 6.3b). This is a v o l u m e w h i c h can be e a s i l y p r o c e s s e d u s i n g only centrifugal ultrafiltration. T h e b a n d i n g p a t t e r n s ( f i n g e r p r i n t s ) reflect t h e d i v e r s i t y of g e n o m e sizes w i t h i n a v i r a l a s s e m b l a g e a n d c o m p a r i s o n s a m o n g s a m p l e s can r e v e a l the s p a t i a l a n d t e m p o r a l scales on w h i c h a s s e m b l a g e c o m p o s i t i o n
98
varies. It should be kept in mind, however, that not all of the viral diversity is necessarily resolved by a one-dimensional separation. Different viruses can have the same g e n o m e size, so band n u m b e r is onlv a m i n i m u m estimate of the total genetic diversity. In addition, much of the viral DNA from environmental samples is clustered in a few narrow size ranges. DNA in these regions appears as somewhat diffuse zones rather than discrete bands. The n u m b e r of different genomes comprising these zones is not known, but could be quite large. Since bands are difficult to resolve in these regions, not all shifts in assemblage composition are necessarily detectable by PFGE alone. However, in practice, spatial and temporal variations in viral c o m m u n i t y composition have been d e m o n strated in a variety of environments (Swain eta[., 1996; Wommack et al., 1999a; Steward and Azam, 2000). Unlike the DNA fingerprinting techniques available for bacteria, viral assemblage fingerprinting by PFGE has the advantage of not involving PCR amplification which can introduce a n u m b e r of biases and artefacts. Since amplification biases are avoided with this method, band intensity is more directly related to the relative abundance of different sized genomes in the original sample. However, several points should be kept in mind. First, measured fluorescence intensity is not necessarily a linear function of DNA mass (Figure 6.4). Deviations from linearitv m a y d e p e n d on the fluorescent stain used as well as the specifications of the system used to record and quantify fluorescence. Therefore, if one wishes to quantify g e n o m e copy numbers, the fluorescent signal must be calibrated using DNA mass standards. Second, dye binding could vary for different viral genomes d e p e n d i n g on factors such as G+C content, presence of modified bases, and whether nucleic acid within the band is single or double stranded, RNA or DNA. To what extent these factors influence the analysis of marine viral assemblages is not well established. It is assmned, however, that most marine viruses contain double-stranded DNA (dsDNA) genomes which should stain with similar efficiencies. Third, there are potential biases in sample preparation. These could include greater losses of larger viruses during 0.2 l~lnl filtration and degradation of more labile viruses during concentration and storage. Again, the influence of these factors is not well known, however, comparison of a filtered and an unfiltered sample indicated that filtration did not dramatically affect the fingerprint (unpublished observation). To minimize the potential for degradation effects, samples should be kept cold and dark, harvested quickly and run on a gel as soon as possible. Another advantage of the PFGE fingerprinting m e t h o d is that the fingerprint is generated using intact viral DNA. This means that for any identified band of interest, the entire viral genome is accessible for further analysis by excising it from the gel. PFGE can thus serve as a starting point for more detailed analyses of viral assemblage composition. This capability is just beginning to be exploited for ecological studies (Wommack et al., 1999b). The detection limit of the assay can be estimated from the dilution series of lambda phage DNA in Figure 6.4. For this gel stained with SYBR Green I (Molecular Probes) and imaged with a laser gel scanner 99
Bacteriophage ~L DNA 0.O1 O . 0 5 0 . 1 9
0.75
3.0
12
ng
(a)
12 (_~ c © u O3 o'3
"E= 0
.._= LL
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E3 >,, t,,,..
6
"13 .I~ I,,._
O3 © ¢...
._Q f,,,_
4 2 O, ) ,
0
, , i , , , i , ~ , i , , , i , , , i , , ,
2
4
6
8
10
12
DNA (ng) Figure 6.4. DNA dilution series showing a nonlinear relationship between DNA mass and fluorescent signal. (a) Digital image of the lambda DNA dilution series acquired by laser gel scanner (Molecular Dynamics). (b) Plot of the fluorescent signal determined as integrated optical density using gel analysis software (RFLPScan; Scanalytics) vs. DNA content of each band.
( M o l e c u l a r D y n a m i c s ) t h e d e t e c t i o n l i m i t w a s _<0.05 n g of D N A in a 5 m m well. A s s u m i n g a 30 k b d s D N A g e n o m e , a b o u t 10" v i r u s e s w o u l d b e d e t e c t a b l e . F o r v i r u s e s w i t h l a r g e r g e n o m e s , t h e d e t e c t i o n l i m i t is e v e n l o w e r , b e i n g a b o u t 10 ~ v i r u s e s for a g e n o m e size of 300 kb. A s s u m i n g t h a t v i r u s e s f r o m 100 ml of s e a w a t e r (i.e. a b o u t 10" v i r u s e s ) a r e l o a d e d in a lane, this m e t h o d is t h e o r e t i c a l l y c a p a b l e of d e t e c t i n g a v i r u s w i t h a 300 kb g e n o m e w h i c h c o m p r i s e s < 0.1% of the c o m m u n i t y . V i r u s e s w i t h a m o r e t y p i c a l g e n o m e of _> 30 k b w o u l d b e t h e o r e t i c a l l y d e t e c t a b l e e v e n if less t h a n 1% of the c o m m u n i t y . T h e p r a c t i c a l d e t e c t i o n l i m i t is l i k e l y to be s o m e w h a t h i g h e r d u e to b a c k g r o u n d f l u o r e s c e n c e in s a m p l e lanes, b u t t h e m e t h o d is c l e a r l y c a p a b l e of d e t e c t i n g e v e n r e l a t i v e l y m i n o r c o m p o n e n t s of the viral a s s e m b l a g e .
I00
eeeeee
CONCLUSION G e n o m i c fingerprinting b y PFGE is a useful tool for exploring the diversity and d y n a m i c s of viruses in the environment. Fingerprints obtained with PFGE can be used to reveal variability in the composition of viral a s s e m b l a g e over space and time. Quantitative analysis of b a n d i n g patterns also p r o v i d e s detailed information a b o u t viral diversity and g e n o m e size distributions in the environment. The ability to access intact viral g e n o m e s from the fingerprints m e a n s that PFGE is also a useful starting point for m o r e detailed analyses. The generality of the m e t h o d m a k e s it a valuable c o m p l e m e n t to the m o r e specific, PCR-based m e t h o d s for analysis of viral diversity and dynamics. Together these techniques p r o v i d e the m e a n s to identify a n d s t u d y the ecology of i m p o r t a n t viral g r o u p s in the ocean regardless of w h e t h e r they can be cultivated. ttl
References
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Birren, B. and Lai, E. (1993). Pulsed Field Gel Eh'ctrophoresis: ,4 Practical Guide, Academic Press, San Diego. Chen, E, Suttle, C. A. and Short, S. M. (1996). Genetic diversity in marine algal virus communities as revealed by sequence analysis of DNA polymerase genes. Appl. Environ. Microbiol. 62, 2869 2874. Fuhrman, J. A. and Suttle, C. A. (1993). Viruses in marine planktonic systems. OceaiTography 6, 51-63. Fuller, N. J., Wilson, W. H., Joint, I. R. and Mann, N. H. (1998). Occurrence of a sequence in marine cyanophages similar to that of T4 g20 and its application to PCR-based detection and quantification techniques. Appl. Environ. Microbiol. 64, 2(151-2060. Klieve, A. V. and Swain, R. A. (1993). Estimation of ruminal bacteriophage numbers by pulsed-field gel electrophoresis and laser densitometrv. Appl. Environ. Microbiol. 59, 2299-2903. Krebs, C. J. (1999). Ecolo2ical Methodology, Benjamin/Cummings, Menlo Park, CA. Sambrook, J., Fritsch, E. F. and Maniatis, T. (1989). Moh'cldar CloIH11~4:A Laboratory Mamlal, Cold Spring Harbor Laboratory Press, Cold Spring }tarbor. Short, S. M. and Suttle, C. A. (1999). Use of polymerase chain reaction and denaturing gradient gel electrophoresis to study diversity in natural virus communities. Hydrobiolo2ia 401, 19-32. Steward, G. F. and Azam, E (2000). Analysis of marine viral assemblages. In: Microbial Biosystems: New Frontiers. Proceedings of the 8th International Symposium on Microbial Ecology (C. R. Bell, M. Brylinski and R JohnsonGreen, Eds). Atlantic Canada Society for Microbial Ecology, Halifax, 159-165. Steward, G. F., Montiel, J. L. and Azam, E (2000). Genome size distributkms indicate variability and similarities among marine viral assemblages from diverse environments. Limmd. Occalmgr. 45, 1697-1706. Suttle, C. A. (1993). Enmneration and isolation of viruses, in: Cum,nt Methods itJ Aquatic Microbiolof~y (P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 121-134. Lewis Publishers, Chelsea. Swain, R. A., Nolan, J. V. and Klieve, A. V. (1996). Natural variability and diurnal fluctuations within the bacteriophage population of the tureen. Appl. E~lvirom Microbiol. 62, 994-997. I01
e- •
° B
Wommack, K. E., Ravel, J., Hill, R. T., Chun, J. and Colwell, R. R. (1999a). Population dynamics of Chesapeake Bay virioplankton: total-community analysis by pulsed-field gel electrophoresis. Appl. Environ. Microbiol. 65, 231-240. Wommack, K. E., Ravel, J., Hill, R. T. and Colwell, R. R. (1999b). Hybridization analysis of Chesapeake Bay virioplankton. Appl. Environ. Microbiol. 65, 241-250.
List of suppliers A/G Technology Corporation 101 Hampton Avenue Needham, M A 02494-2628, USA Teh +1 781 449 5774 or +1 800 248 2535 Fax: +1 781 449 5786 E-maih
[email protected] Internet: www.agtech.com
Tangential flow ultrafiltration products Amersham Pharmacia Biotech AB SE-751 84 Uppsala, Sweden Teh +46 (0) 18 612 O0 O0 Fax: +46 (0) 18 612 12 O0 lnternet: www.apbiotech.com
Gel documentation/fluorescence DNA quantification system (Molecular Dynamics FluorImager), fluorometers Bio-Rad Laboratories Life Science Research Group 2000 Alfred Nobel Drive Hercules, California 94547, USA Tel: +1 800 424 6723 Fax: +1 510 741 5800 Internet: www.bio-rad.com
Pulsed field gel electrophoresis systems and supplies, Gel documentation systems, fluorescence DNA quantification equipment and reagents
BioWhittaker Molecular Applications 191 Thomaston Street Rockland, ME 04841, USA Internet: www.bioproducts.com Teh +1 800 341 1574 Fax: +1 207 594-3426
Agarose Fisher Scientific 2000 Park Lane Pittsbuq~h, PA 15275, USA Teh +1 800 766 7000 Fax: +1 800 926 1166 lnternet: wwwl.fishersci.com
Centrifuges, general laboratory supplies and equipment, plasticware, chemicals, Polaroid film GIBCO/BRL Life Technologies, Inc. 9800 Medical Center Drive Post Office Box 6482 Rockville, M D 20849-6482, USA Teh +1 800 338 5772 Fax: +1 800 3312286 Internet: www.lifetech.com
DNA mass ladder Kendro Laboratory Products 31 Pecks Lane NeWtown, CT 06470-2337, USA Teh +1 800 522 7746 Fax: +1 203 270 2166, +1 203 270 2210 or +1 203 270-2115 E-maih in[
[email protected] Internet: www.sorvall.com
Centrifuges and rotors
102
Millipore, Inc.
Phoretix International
80 Ashby Road Bedford, MA 01730-2271, USA Tel: +1 781 533 6000 Fax: +1 781 533 3110 Internet: www.millipore.com
Tyne House, 26 Side Newcastle upon Tl/ne NE1 3JA, UK Tel: +44 (0)191 230 2121 Fax: +44 (0)191 230 2131 E-maih
[email protected] hzternet: www.phoretix.com
Tangential flow and centrifugal ultrafiltration products (Millipore and Amicon brands), membrane filters
Gel analysis software
Scanalytics, Inc. Molecular Probes, Inc. 4849 Pitchford Ave. Eugene, OR 97402-9165, USA Teh +1 541 465 8300 Fax: +1 541 344 6504 E-mail:
[email protected] [tlteFitet" ww~t~.]ll'ob?s,COltl
Fluorescent nucleic acid stains (SYBR Green I, PicoGreen)
8550 Lee Highway, Suite 400 Fairfax, VA 22031-1515, USA Tel: +1 703 208 2230 Fax: +1 703 208 1960 E-maih
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Gel analysis software
New England BioLabs
103
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Enzymes, chemicals, biochemicals, Polaroid film, DNA quantification standards and DNA size markers
DNA size markers
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Sigma-Aldrich 3050 Spruce Street St. Louis, MO 63103, USA Trl: +1 314 771 5765 or +1 800 325 8070 Fax: +1 314 771 5757 E-maih
[email protected] lnternet: www.sigma-aldrich.com
32 Tozer Road Beverly, MA 01915, USA Teh +1 800 632 5227 Fax: +1 800 632 7440 E-mail:
[email protected] hlternet: ~O~U'Ud.Ileb.co?tI or wWW.ltk.tleb.coltl
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7 Lysogeny and Transduction John H Paul' and Sunny C Jiang2 ~Department of Marine Science, University of South Florida, St. Petersburg, FL 33 70 I, USA 2School of Social Ecology, University of California, Irvine, Irvine, CA 92697, USA
CONTENTS Introduction and background Screening marine bacteria for lysogeny Transduction assay Future directions
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# INTRODUCTION
AND BACKGROUND
Lysogeny and transduction describe a type of phage/host interaction and a method of bacterial gene transfer (procaryotic sex), respectively. Although they are often reviewed together, these topics are linked only in that one type of transduction (specialized) has an obligate requirement for a lysogenic interaction. In this chapter we describe the background for understanding both of these processes, and give methods that we have found useful in studying lysogeny and transduction in the marine environment.
Lysogeny and pseudolysogeny Lysogeny occurs when a phage enters into a stable symbiosis with its host (Ackermann and DuBow, 1987). The host (bacterium or algal cell) and phage capable of entering into such a relationship are termed a lysogen and temperate phage, respectively. The temperate phage genome becomes integrated into one of the replicons of the cell (chromosome, plasmid, or another temperate phage genome) and is termed a prophage (Figure 7.1). The lysogenic state is a highly evolved state (Levin and Lenski, 1983) requiring coordinated expression (and repression) of both host and viral genes. There is a selective pressure to favor lysogeny, particularly at times of low host density, because a temperate phage is less likely to drive its host to extinction (Levin and Lenski, 1983). Other advantages of lysogeny include the expression of prophage encoded genes, termed conversion. This is in contrast to transduction (see below), whereby the genes imparted into an infected host were the result of a phage packaging error during a prior infection cycle. That is, in transduction the genes METHODS IN MICROBIOLOGY, VO1,UME 30 ISBN 0-12-521530 4
C o p y r i g h t © 200l A c a d e m i c Press Ltd All rights of r e p r o d u c t i o n in any form reserved
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Lytic
Pseudolysogenic
Lysogenic
Figure 7.1. Cartoon depicting lytic, lysogenic, and pseudolysogenic phage/bacterial interactions. The oval ring in the bacterial cell is the bacterial chrocnosome. After phage adsorption, the phage DNA is injected into the bacterium, and is represented as a coiled molecule in the figure. This can become integrated into the host chromosome as a prophage in lysogeny (indicated as a straight bar in the bacterial chromosome in the right side of the figure). By the process of induction, the prophage is excised, and can go into lytic phage replication (left side of the figure). In pseudolysogeny, the host cell can mutate to an adhesion-impaired or deficient state (depicted by a wavy cell surface), whereby collisions result in a low success rate of infection. Another type of pseudolysogeny (termed carrier state) can occur when the prophage does not integrate but is maintained as a plasmid (central panel in the figure). Both types of pseudolysogeny result in a high abundance of both phages and host cells simultaneously.
originated in a bacterial host and are not a normal part of the p h a g e genome. Traits conferred to the host by conversion include i m m u n i t y to superinfection, which m e a n s that the host becomes i m m u n e not only to infection by that particular t e m p e r a t e phage, but to other closely related phages. Other traits include antibiotic resistance, restriction modification systems, and toxin a n d / o r bacterial virulence (as for diptheria toxin and botulinus toxins C and D; A c k e r m a n n and DuBow, 1987). In a lysogenic symbiosis, there is a low rate of s p o n t a n e o u s reversion to virulence, in that, on average, 1 in 10 ~ produces a lyric event ( A c k e r m a n n and DuBow, 1987). Thus, there is a constant production of t e m p e r a t e p h a g e in a lysogenic interaction. However, other interactions exist where there is a high-level p r o d u c t i o n of both host cells and viral particles in a process termed ' p s e u d o l y s o g e n y ' (Figure 7.1). P s e u d o l y s o g e n y has been used s y n o n y m o u s l y with 'carrier state' and 'chronic infection', yet distinctions can be m a d e between these conditions. By A c k e r m a n n and D u B o w ' s (1987) definition, p s e u d o l y s o g e n y is a p h e n o m e n o n caused by a mixture of sensitive and resistant host cells, or a mixture of t e m p e r a t e and virulent
106
phage, that results in a constant supply of host and viral particles. This in many ways resembles the marine environment, where sensitive and resistant cells coexist with lyric and temperate phages of many differing strains and species. We have indicated two such interactions that may occur in pseudolysogeny in Figure 7.1. The first is a mutation to an adhesion-impaired or deficient state, thereby limiting the number of successful infections. Also shown is what has been termed the carrier state; a pseudolysogenic-like relationship occurs characterized by plasrnid-like prophages, which do not integrate into the host genome (Figure 7.1). Chronic infection is the process whereby certain bacteria produce phage without host lysis, by budding or extrusion, as in Pl or M13 (Dehardt et al., 1978). In true lysogeny, when a temperate phage infects a host, a 'lysogenic decision' is made, as to whether a lytic or lysogenic interaction will ensue. Factors affecting the lysogenic decision are the multiplicity of infection (MOI; a high MOI favors lysogeny), host growth rate, and nutrient status (Levin and Lenski, 1983). In fact, Wilson and co-workers (Scanlan and Wilson, 1999; Wilson et al., 1998) have hypothesized that phosphate concentrations influenced the lysogenic decision in cyanophage infecting Synechococcus. When phosphate-limited microcosms containing a bloom of Synechococcus were enriched with Pi (inorganic phosphate), there was a dramatic increase in phage production concomitant with a crash of the Synechococcus population (Wilson et al., 1998). Lysogeny in Synechococcus populations would be consistent with the observation of high cyanophage abundance yet resistance to infection (Waterbury and Valois, 1993). Lysogeny is extremely common amongst bacteria, at least in cultivated strains. Ackerman and DuBow (1987) indicated that among 1200 diverse strains of bacteria, an average of 47~7, contained inducible prophage. Jiang and Paul indicated that among 110 marine bacterial isolates, 40(7,, were lysogenized (Jiang and Paul, 1998a). The importance of lysogeny among natural populations of bacteria is a topic of debate. Wilcox and Fuhrman (1994) concluded that lytic infection was far more important than lysogeny in bacterial mortality or phage production based upon studies with natural populations exposed to sunlight. Weinbauer and Suttle (1996, 1999) also concluded that a small proportion (1.5-11.4%) of the bacteria in marine samples from the Gulf of Mexico were lysogenized, with the highest values occurring for offshore populations. Tapper and Hicks (1997) estimated from 0.1 to 7.4% of the bacteria in Lake Superior to be lysogens, in agreement with other studies. Our lab has studied the distribution of lysogeny in various environments, and found eight of ten eutrophic estuarine environments to contain inducible prophage, whereas only three of eleven offshore environments were positive for prophage induction. We have shown that a series of environmentally relevant pollutants (polynuclear aromatic hydrocarbons, polychlorinated biphenyls, and pesticides) can all cause induction of natural populations of lysogens (Jiang and Paul, 1996; Cochran et al., 1998), and that there was a seasonality in the detection of lysogeny, with lysogens prevalent in the summer months, but absent in winter (November to February; Cochran and Paul, 1998). Our estimates of the 107
percentage of bacteria lysogenized are in agreement with others, ranging from undetectable to 37%, averaging 6.9%, based upon an assumed burst size of 30. If 8% of the population was lysogenized, and half of these were induced by some environmental factor, this would produce 5 x 10'~ phage ml ', or nearly half of the phage present in Tampa Bay. If nutrients and temperature can control prophage induction a n d / o r the lysogenic decision, it seems reasonable that induced temperate phage may constitute a significant amount, or perhaps the majority, of phage present in many coastal environments. The detection of lysogeny in cultures or natural populations is usually through prophage induction by use of a mutagenic agent, usually mitomycin C. The methods described below are all based on some derivative of this procedure.
Gene transfer by transduction Bacteriophage-mediated transduction is one of three well-known mechanisms, along with conjugation and transformation, of horizontal gene transfer among prokaryotic organisms. In transduction, bacterial DNA or plasmid DNA is encapsulated into phage particles during lytic replication of the phage in the donor cell and is transferred to the recipient cell by infection. This donor DNA either undergoes recombination with the host chromosome to produce a stable transductant or remains extrachromosomal as a plasmid. Based on the mechanism of production of transducing viral particles and the means by which DNA is incorporated into the recipient cell chromosome, transduction can be classified as one of two types, specialized or generalized. In specialized transduction, only a restricted number of genes within tile host may be transferred; namely, those which flank the site of integration of the prophage. Specialized transducing particles are produced only from the integrated prophage during induction (Buck and Groman, 1981; Cavenagh and Miller, 1985). The DNA present in transducing phage particles is produced by aberrant excision of prophage DNA (Sternberg and Maurer, 1991). When this DNA is injected into the recipient host during infection, it is established and maintained as a prophage independent of the host's general homologous recombination system. In contrast, all regions of the chromosome or other genetic elements present in the donor cell can be transferred with approximately the same frequency in generalized transduction (Zinder and Lederberg, 1952). Following injection of this DNA into the recipient cell, stable transductants are generated by the replacement of the homologous cell DNA with the transducing DNA. Therefore, generalized transduction is dependent on the general recombination system of tile host (Sternberg and Maurer, 1991). Virulent as well as temperate bacteriophages are capable of generalized transduction under appropriate conditions. Genomics studies have indicated that considerable horizontal gene transfer has occurred between prokaryotes (Jain et al., 1999). The transfer of genetic information between distantly or even unrelated organisms
108
during evolution has been inferred from nucleotide sequence comparisons (Dr6ge et al., 1998). Among the bacterial gene exchange mechanisms, transformation and conjugation were identified as mechanisms with potentially the broadest host range of transfer. However, sufficient evidence has accumulated to indicate that transduction is a significant mechanism of gene transfer, being more important in natural ecosystems than originally thought (Novick et al., 1986; Kokjohn, 1989; Saye and Miller, 1989; Stozky, 1989; Saye et al., 1990; Miller et al., 1992; Schichlmaier and Schmierger, 1995). Because the packaging of nucleic acids in a phage particle may represent an evolutionary survival strategy for the genetic material, bacteriophages may serve as reservoirs for exogenous genes (Zeph et al., 1988). Transduction has now been shown to be an important mechanism of gene transfer within several natural ecosystems, including soils (Germida and Khachatourians, 1988; Zeph ct al., 1988; Stotzky, 1989; Zeph and Stotzky, 1989), plant surfaces (Kidambi et al., 1993), freshwater environments (Morrison et al., 1978; Saye et al., 1987; 1990; Amin and Day, 1988; Miller, 1992; Ripp et al., 1994; Ripp and Miller, 1995) and animals (Jarolmen et al., 1965; Novick and Morse, 1967; Baross et al., 1978; Novick et al., 1986). Both chromosome and plasmid transduction in Pseudomonas aeruginosa were demonstrated during in situ incubation in a freshwater lake (Morrison et al., 1978; Saye et al., 1987; 1990) and on submerged river stones (Amin and Day, 1988), with transduction frequencies ranging from 1.4 x 10 ~to 8.3 × 10 ~/recipient. Ripp and Miller (1995) also suggested that the presence of suspended particulates in the water column facilitates transduction by bringing the host and phage into close contact with each other. Compared with freshwater environments, less is known about transduction in marine waters even though a transducing marine bacteriophage was isolated more than 15 years ago (Keynan et al., 1974). Over the past ten years, bacteriophages were found to be the most numerous microorganisms in the ocean. In addition, bacteriophages may have a broader host range than previously expected. Jensen et al. (1998) have demonstrated the prevalence of broad-host-range lyric bacteriophages (90%) in both a freshwater pond and sewage waters. They also suggest that standard bacteriophage enrichment using a single bacterial host is unavoidably biased against the development of viruses with a broad host range, and this bias may partially explain the general view that bacteriophages are restricted in their interactive host range. Wichels et al. (1999) found that 8% of 62 marine bacteriophage isolates examined were capable of infecting a variety of hosts. The host ranges consist of 11 to 36 unique bacterial isolates. The prevalence of broad-host-range lytic bacteriophages has profound ecological significance, especially with regard to natural mechanisms for gene transfer. Jiang and Paul (1998b) described a plasmid transduction system using a temperate marine virus and host isolate (Figure 7.2). Transfer of an antibiotic resistant plasmid by this phage was detected at a frequency of 10 ~-10" per pfu (plaque forming unit). Interestingly, all transductants were also lysogenized with the temperate phage genome. To investigate 109
co
e-
# e-
¢,.
0
H©PE -1 (~HSIC Kan + Strep
Treatment
Lysate
+@
Plasmid Hybridization
Kan + Strep
Control Figure 7.2. Cartoon depicting a plasmid transduction experiment using a plasmid containing donor (indicated as the HOPE-1 bacterium; Jiang and Paul, 1998b) and the phage 0HSIC. A lysate is made from the plasmid-containing host and used to infect the wild-type host. Survivors are plated on media containing antibiotics, the resistances for which were encoded in the transducing plasmid. The appearance of antibiotic-resistant bacteria that hybridize to a probe for the transducing DNA indicates that transduction has occurred. A no-Iysate control is included, which yields no colonies, and does not hybridize to the gent probe specific for the plasmid.
transduction to the indigenous marine bacterial community, Jiang and Paul (1998b) used the concentrated marine bacterial c o m m u n i t y from various environments as recipients (Figure 7.3). Transduction was found in two sampling sites at a frequency of 10 ~ per pfu. The transductants were confirmed by PCR amplification of plasmid-specific sequences. Chiura (1997) reported the first intergeneric phage-mediated g e n t transfer between marine bacteria and enteric bacteria. He demonstrated that five marine bacteria isolated from seawater were capable of spontaneous induction of temperate phages after a prolonged incubation. Although these phages did not form plaques on an E. coli bacterial lawn, they were capable of generalized transduction of genes to repair amino acid deficiencies in E. coll. The five marine isolates were not phylogenetically closely related and none of them were closely related to E. coll. Auxotrophic markers on the E. coli c h r o m o s o m e exhibited gene transfer frequencies ranging between 10 ~and 10 ~ per viral particle. Therefore, the transducing frequencies of these viral particles spontaneously induced from marine bacteria, were four to seven orders of m a g n i t u d e higher than those of transducing phages isolated from freshwaters (Saye et el., 1990; Ripp et el., 1994). Intergeneric transduction was also demonstrated in another startling report in which viral-like particles (VLP) produced by a hot-spring natural bacterial c o m m u n i t y (predominated by hyperthermophilic chemolithotrophic sulfur bacteria) were capable of transducing loci
II0
to repair auxotrophic E. coli and Bacillus subtilis to prototrophy with an average efficiency of 10 ~'per VLP (Chiura et al., 1998). These results indicate that spontaneous viral production by marine bacteria may be an important mechanism of generalized horizontal gene transfer involving a broad range of bacterial hosts in the marine environment.
4,e4,ee4, S C R E E N I N G M A R I N E B A C T E R I A F O R LYSOGENY Isolates in c u l t u r e The protocol that follows has been used to screen marine bacterial isolates for inducible prophage a n d / o r bacteriocin-like particles (Jiang and Paul, 1994; 1998a). The protocol was developed for rapidly growing cultures in flasks but has been readily adapted to microtiter plates. We have used it only with our formulation of marine bacterial growth medium (ASWJP+PY; Paul and Myers, 1982) but any heterotrophic bacterial medium should work equally as well. Bacteria are grown into exponential phase in batch culture and then exposed to mitomycin C (or another mutagen such as UV light). The growth of the culture is followed by optical density (absorbance) and prophage induction is detected by a decrease or stasis in absorbance compared to a control (unamended) culture. Viral counts are made (either by TEM or epifluorescence microscopy) for both treated and control cultures. A significant increase in viral particles over the control is indicative of lysogeny.
Materials and supplies • • • • •
Marine bacterial isolate(s) as frozen glycerol stock or from agar plate (<1 month old) ASWJP+PY medium (Paul and Myers, 1982), Zobell's 2216 Marine Media (Difco), or other marine bacterial medium 0.02qam-flltered DI water 0.02-1urn-filtered formalin All materials for SYBR Green or SYBR Gold viral direct counts (see Chapter 3 by Noble)
Assay 1.
2.
Grow the selected culture of marine bacteria overnight in rich medium. This can be accomplished by using 0.5 ml of a frozen stock in 5 ml media in a sterile 15 cc centrifuge with shaking (200 rpm). In the morning, add 0.5-2.0 ml of the culture to 50 ml sterile marine bacterial media, incubating at the correct temperature, again with shaking.
III
3. 4.
5.
6.
Take 1 ml samples every hour and monitor absorbance at 600 nm. When absorbance reaches between 0.4 and 0.6, immediately take a 1.0 ml sample and centrifuge in a microcentrifuge for 5 min at 14 000 rpm. Remove 0.5 ml of the supernatant, add it to 4.5 ml of the 0.02-Mm-filtered DI water and 125 ~1 of the 0.02-~tm-filtered formalin (final concentration -1%). This can be stained directly for SYBR Green or Gold viral counts (500 B1 aliquots or diluted an additional 1:10 with 0.02-~tm-filtered DI for counting). Split the culture in two equal aliquots (20 ml each). Add Mitomycin C to one (final concentration 0.5 Mg ml ~). Continue incubating with shaking and take an absorbance reading every hour. Sample again at 3 h, 8 h (optional) and overnight (-16 h) for viral counts as in step 4. Compare viral counts in the Mitomycin C treatment to the t = 0 and control at equal time points.
Screening for lysogens in natural populations Questions that arise often in the study of lysogeny in the marine environment include 'How common an event is lysogeny? What proportion of the bacterial population contain inducible prophage (are lysogens)? What proportion of the viral population is temperate, or is the result of a prophage induction event?' These questions cannot be unequivocally answered with current technology. Using the methods below, it is possible to determine how common lysogeny is when comparing different marine or estuarine environments or different times of the year at the same environment. Making a few assumptions, it is possible to estimate the proportion of the bacteria which contain inducible prophage (and therefore are by definition lysogenic). With speculation, one might be able to arrive at an estimate of the proportion of the phage population which is the result of prophage induction events. The techniques described below have been successfully used with natural populations to detect the occurrence of lysogeny (Jiang and Paul, 1994; Cochran and Paul, 1998), or to determine if certain environmental pollutants can cause prophage induction by natural populations (Jiang and Paul, 1996; Cochran et al., 1998). The inducing agent has been Mitomycin C unless other pollutants were investigated. We have examined the capability of natural populations to be induced by mutagens either with samples concentrated by Membrex ultrafiltration (Jiang and Paul, 1996) or with unconcentrated natural populations. The former method has been employed in oligotrophic, offshore environments where bacterial populations were well less than 10" ml '. However, use of Membrex concentration to determine tile number of lysogens present is probably not quantitative because of the potential for bacterial growth during the concentration process. Both methods have been used in conjunction with TEM for viral enumeration, although we have almost exclusively changed to SYBR Gold enumeration using epifluorescence microscopy.
112
Equipment and supplies •
• • •
•
Membrex Rotary Biofiltration Device, equipped with 100 KD filter (note that this is necessary only for offshore samples and use withTEM enumeration Mitomycin C (Sigma) or other mutagens to be tested Sterile 15 or 60 ml conical centrifuge tubes TEM-grade glutaraldehyde (forTEM enumeration only) 0.02-pm-filtered formalin (for epifluorescence viral enumeration only)
Assay 1.
2.
3.
4.
5.
6.
7.
For Membrex concentration of the ambient microbial populations, 10-100 1 of sample are concentrated using the rotary biofiltration device as described in the manufacturer's instructions. The concentrate (termed retentate) usually has a v o l u m e of 35-60 ml. For concentrated samples, place 1 ml in a sterile 1.5 ml microcentrifuge tube or 5.0 ml of the retentate in a 15 ml conical centrifuge tube. Do this both for control (unamended) and treatment (mutagen a m e n d e d ) samples. For unconcentrated samples, add 25 ml each to a control or treatment, 50 ml sterile, conical centrifuge tubes. If several mutagens are to be investigated, increase the n u m b e r of treatment tubes accordingly. Take an additional sample (1-5 ml for Membrex Retentate, 25 ml for unconcentrated) and fix with glutaraldehyde (2% final concentration) as a T = 0 control. If epifluorescence microscopy is to be used for enumeration, fix sample with 1% 0.02-~tm-filtered formalin. For the treatment samples, add 0.5-1 Hg ml ' Mitomycin C. If other mutagens are to be used, it is a good idea to include a Mitomycin C treatment as a positive control. Mutagens can be added at any concentration desired, but this can be limited by the solubility of the m u t a g e n (e.g. Polynuclear aromatic hydrocarbons; Jiang and Paul, 1996). The samples are incubated for 16-24 h at room temperature and either fixed with 2% glutaraldehyde (for TEM) or 1°/, formalin (epifluorescence microscopy). Samples for enumeration by epifluorescence microscopy should be counted within 24 h of collection. For TEM samples, if the sample has not been concentrated by Membrex ultrafiltration, use ultracentrifugation (160000g) to impinge viral particles onto Formvar-coated TEM grids (Borsheim et al., 1990). If the samples have been concentrated by ultrafiltration, it may be necessary to dilute the sample 1:10 with DI water before spotting 1 H1 onto a Formvar grid. Count both bacteria and viruses in control and treated samples. For induction to have occurred, viral counts in the treatment must exceed those in the control.
113
e,. O
u "o e-
# "0 >., C ~0 0
8.
Calculate the percentage lysogenic bacteria as follows: % lysogens = [(VDC, - VDC¢)/B~]/BDC,
,~
where VDC: is the viral direct counts (in viruses ml ') in the treatment, VDCc is the viral direct counts in the control, B~ is the average burst size, and BDC, ,, is the bacterial counts at the set up of the experiment (T = 0). The average burst size can be derived by TEM observation of bacterial bursts. We have found an average for our samples from the Gulf of Mexico of 30, whereas taking an average of the literature from a recent review (Wommack and Colwell, 2000) indicates a value of 53.5 _+48.
P r o p h a g e i n d u c t i o n in n a t u r a l p o p u l a t i o n s m v i r a l r e d u c e d m e t h o d The m e t h o d described above for detection of lysogeny in natural populations has the least a m o u n t of manipulation of the sample. However, the ambient levels of viruses will confound detection of small increases in viral counts because of prophage induction. To obviate this problem, Weinbauer and Suttle (1996) used a technique to reduce the level of ambient viruses by filtration of the ambient c o m m u n i t y through a 0.2 btm filter and washing the c o m m u n i t y in viral-free water. In a seasonal study of lysogeny currently u n d e r w a y in our laboratory, this procedure reduced viral direct counts by 62% while decreasing bacterial direct counts by 35%. In this study over five samplings, prophage induction was detected only by the viral reduced method.
Materials and supplies All items listed in the section on Screening for lysogens in natural populations • Sterile 47 mm polycarbonate filtration devices with reservoirs •
• • •
Sterile 47 mm Anodisc filters, 0.02 ~tm Sterile 47 rnm Nuclepore or Poretics filters, 0.2 ~m Sterile 47 mm Nuclepore or Poretics filters, 1.0 pm
Protocol 1.
2.
The water sample (300 to 1500 ml) is first filtered through a 1 ~m filter to remove protozoan grazers. We often omit this step in estuarine waters because of the n u m b e r of bacteria which are greater than 1 gm in size. Prepare 0.02-btm-filtered water using one of the sterile polycarbonate filtration devices and the 47 m m 0.02 btm Anodisc filters.
114
3.
4.
5.
6. 7.
Marine
Set up a second sterile polycarbonate filtration device with a 47 m m 0.2 btm filter and gently filter the water sample (we typically use 60 ml), turning off the v a c u u m w h e n the v o l u m e is reduced to about 5 ml, making sure not to filter to dryness. A d d 40 ml of the virus-free sample water to the u p p e r reservoir of the filtration device containing the 5 ml of filter-concentrated sample. Again filter until the v o l u m e is reduced to 5.0 ml. Using a sterile 10 ml pipette, collect the concentrated water sample and place it into a sterile 125 ml p o l y m e t h y l p e n t e n e flask. Using sterile forceps, remove the 0.02 btm filter and add it to the flask, along with 40 ml of additional 0.02-btm-filtered water. Vortex for 30 s, then remove the filter with sterile forceps. Bring the v o l u m e to 60 ml with 0.02-~m-filtered sample water. At this point we typically fix 10 ml for T -- 0 viral and bacterial counts, and use 25 ml each for treatment (i.e. Mitomycin C) and control prophage induction assay. Samples are then counted as described in the protocol above.
prophage
induction
assay
This protocol uses cultures of marine bacteria in an assay that can either be used to detect mutagenic activity of samples or c o m p o u n d s w h e n using a k n o w n lysogen, or used to detect lysogeny in marine bacterial isolates. It is a rapid way of also detecting the sensitivity of the bacteria to mutagens because it uses a range of concentration of mutagen. A disadvantage is that the level of induction m a y not be as great in the microtiter plate format because of limited aeration compared to rapidly shaking, well-aerated flasks.
Materials and supplies • 96-well microtiter plate with lid (i.e. Costar 3799 96-well Cell Culture Cluster) • Stock solutions of mitomycin C: 2.5~g ml' and 0.5~g ml' in Marine Nutrient Broth (ASWJP+PY) • Marine Nutrient Broth (ASWJP+PY) • Overnight marine bacterial culture
Assay 1.
Depending on the n u m b e r of bacteria to be assayed, designate four rows of the microtiter plate per strain, two for Mitomycin C, two for control.
115
2.
3.
4.
5.
6.
7. 8. 9.
,,~,,t
Inoculate a fresh flask (i.e. 10-25 ml) with the overnight culture. Monitor growth as A~,~,,,and w h e n the absorbance reaches 0.4-0.6, use the cells in the assay. Determine which rows are to be used for Mitomycin C, and add 55 ~tl of 2.5 gg ml ~Mitomycin C to the first well in those rows, and 55 gl of 0.5 ~g ml ' to the second well in those rows. Add 55 ~i of Marine nutrient broth to the first and second wells in the control rows. For the u n k n o w n (treatment rows) add 55 gl of the appropriate u n k n o w n sample to the first well, and then 55 gl of a 1:5 dilution of the u n k n o w n to the second row. Using an Octapipette, pipette 50 gl of nutrient broth into all the other wells. Note: it is probably necessary to go to only six or eight columns (final conc. 0.5-1 ng ml '). Using a multichannel pipettor (i.e. Octapipette) set for 5 ~1, transfer 5 ~1 from the first column to the third column for a 1:10 dilution. Triturate to mix. Then proceed to the fifth and the seventh, if necessary, performing 1:10 dilutions with trituration. Using an Octapipette set for 5 gl, transfer 5 gl from the second column to the fourth column, for a 1:10 dilution. Triturate to mix. Then proceed to the sixth and repeat. This will result in a dilution series starting with 0.5 gg ml ', and including 0.1, 0.05, 0.01, etc. until 0.001 at the sixth column. A d d 200 gl of the exponentially growing cells to all wells. A d d the lid to the microtiter plate and rock very gently (no sloshing) overnight (16 h) at the correct temperature for growth. At the end of the experiment, add 6.7 gl of 0.02-~tm-filtered formalin to each well. Pipette the contents of the wells into microcentrifuge tubes. Centrifuge the bacteria at 14 000 rpm in a microcentrifuge for 5 rain. Collect 200 gl of the supernatant and dilute appropriately for SYBR Gold counts. Positive induction is determined as a significant increase in viral counts over controls.
TRANSDUCTION
ASSAY
Transduction c o m p o n e n t s and conditions Three basic components are required for a transduction system: a donor, transducing phage and a recipient. In most transduction assays, both the d o n o r and recipient should be sensitive to infection by the same bacteriophage. Both cell-free phage lysates resulting from lyric phage production and phage particles produced by spontaneous induction of lysogens can mediate transduction. Also, both lysogenic and non-lysogenic bacteria can serve as recipients, but lysogenic recipients have higher transduction frequencies, possibly due to the lysogenic protection (homoimmunity) from lytic attack (Miller et al., 1992).
116
In theory, all bacteriophages are capable of generalized transduction at various frequencies because mistakes in packaging of DNA within the bacterial host always occur (Ackermann and DuBow, 1987). However, to effectively detect or demonstrate the process of transducfion, phenotypical or genotypical markers are necessary to monitor the acquisition and expression of transduced genes. Auxotrophic mutants with specific amino acid requirements for growth or plasmids encoding for antibiotic resistance are often the biomarker of choice. Both types of markers allow selection of transductants from recipients by plating on selective medium and therefore reducing the background growth of recipient bacteria. Proper control experiments are critical to subtract the rate of spontaneous revertants from transduction (Levisohn et al., 1987). Cotransduction of closely linked loci will allow a more definitive identification of a unique transduced phenotype and reduce the background of revertants produced by spontaneous mutation (Miller, 1992). Compared to transduction of chromosomal markers for which a good gene probe does not exist, transduction of antibiotic resistant plasmids is more easily confirmed, either by plasmid DNA extraction from transductants followed by restriction enzyme profile analysis (Ripp et al., 1994), or by colony hybridization with specific gene probes (Jiang and Paul, 1998b; Figure 2). Control experiments are also necessary to correct for rates of spontaneous mutation to resistance. If it is necessary to further confirm the transfer event, plasmid extraction, Southern hybridization or PCR techniques can also be used to verify the existence of the original plasmid in the transductants. However, rearrangement or recombination of the plasmid DNA can occur, particularly when natural populations are employed as recipients (Jiang and Paul, 1998b). Another problem in the use of antibiotic resistance plasmids for use in transfer to indigenous recipients is the high degree of antibiotic resistance found in natural populations. In our studies of transfer of the plasmid pQSRS0, which encodes kanamycin resistance on the transposon Tn5 as well as streptomycin, a high level of resistance was often found to both kanamycin and streptomycin in marine bacterial populations. Additionally, some of the resistant colonies in the 'no plasmid control' hybridized with a gene probe derived from the kan resistance gene of Tn5 (Figure 7.3). When such results were obtained, the results were discarded, and only environments lacking Tn5-1ike kanamycin resistance were studied further (Jiang and Paul, 1998b). The size of the plasmid used in a transduction assay should be considered. Saye et al. (1987) found that transduction of plasmids was more efficient if the molecular weight of the plasmid was similar to that of the phage genome, favoring packaging errors. The frequency of transduction varies with different phage-host systems. However, exposing transducing particles to UV radiation is generally known to increase transduction frequency (Miller, 1992). It has been suggested that this treatment stimulates recombination within the host cell leading to increased incorporation of the transduced DNA into the recipient genome (Benzinger and Hartman, 1962). Secondly, it may 117
HOPE -2 (1)DIB Kan + Strep
Plasmid Hybridization
}.
-I-
)
or
Treatment
Lysate Kan + Strep
+ Control
Figure 7.3. Cartoon depicting plasmid transduction as in Figure 2 but using the ambient microbial population (depicted as a tank containing copedods and fish) as recipients. Unlike transduction with a known recipient, there is an indigenous level of antibiotic resistance in the natural population, which yields colonies from the no lysate control when plated upon kanamycin/streptomycin media. In certain cases, some of these hybridize to the probe, showing that the indigenous population contains some genes similar to those chosen for transduction. Only when tile no-lysate controls contain no positive hybridizing colonies can transduction be inferred.
reduce the infectivity (or virulence) of the phage, such that all putative transductants are not lysed as the result of lytic infections. The MOI (multiplicity of infection) is another factor influencing the frequency of transduction (Keynan et al., 1974; Morgan, 1979). In general, the optimal MOI range is between 0.1 and 1. It is thought that low MOIs produce higher transduction frequencies by reducing the possibility of a recipient cell simultaneously encountering both a transducing particle and a lytic infectious phage particle (Miller, 1992). For transduction in environmental chambers, Saye et al. (1990) reported optimal MOIs for phage Fl16 and DS1 transduction in P. aeruginosa to be 0.02. Jiang and Paul (1998b) found marine transduction occurred only w h e n the MOI was less than 0.05.
Transduction
in c u l t u r e d isolates
To assay transduction in a cultured p h a g e - h o s t system, the following steps can be followed to establish a transduction system. However, the m e t h o d presented here is highly generalized and can be adapted to various plasmids and p h a g e - h o s t systems.
118
Materials and supplies • • • • • • • •
• • • • • •
Marine bacteria and bacteriophage isolates Plasmids (preferably broad host range with two selectable markers and accompanying gene probes) Antibiotics Marine broth and other nutrient medium (Difco) Bacto agar (Difco) Petri dishes Culture tubes 0.5 MTris.HCI buffer pH 8.0 N-methyI-N'-nitro-N-nitrosoguanidine (Sigma) 0.2 lure membranes (Millipore) Chloroform E~Nase I (Sigma) UV lamp ~/ater bath
cO U "O e-
#
Methods
C
Construction of gen etically marked donor The protocol below describes transferring an antibiotic resistant plasmid to a donor strain by triparental mating. Alternative methods of plasmid transier, i.e. artificial transformation, can also be used to achieve the same goal. Protocol
1. Mix log-phase cultures of the following strains, a plasmid donor, a helper strain containing conjugative helper plasmid and the plasmid recipient, at approximately equal cell numbers. 2. Fi ter the mixture onto a sterile, 0.2 gm membrane filter and incubate ox ernight on nonselective medium to allow conjugation. 3. The next morning, re-suspend cells in 5 ml of nutrient medium, then plate onto selective medium to select for the traits of plasmid and recipients.
Production of transducing phage particles from a lytic infection of donor cells Protocol
1. Mix midlog donor bacteria with phages at a MOI of -0.1 in 3 ml of melted 1% agar medium kept at 45-47°C in a water bath. 2. Pour the soft agar mixture over a 1.5% agar plate containing the proper growth medium. For some marine phage-host systems that are sensitive to a brief exposure to higher than ambient temperature, the soft agar should be taken out of the water bath just before adding the phage-host mixture then poured immediately to prevent heat inactivation. 119
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3. Incubate overnight for phage amplification, harvest phages by flooding the top agar with 0.5 M Tris buffer (EH 8.0), using 5 ml for each 110 mm diameter plate. 4. Remove cell debris or residual bacteria by low speed centrifugation followed by filtration through a sterile 0.2 ~m filter. Alternatively, a drop of chloroform is added to kill residual bacteria. 5. Repeat steps 1 to 4 for a second round of infection to ensure that the transducing lysate contains markers derived from the donor host. 6. If desired, treat transducing lysates by ultraviolet radiation using a lamp with a peak wavelength at 256 nm to reduce the infective phage titer to 1% of the original. 7. Digest transducing lysates with 50 units ml ~of DNase I before use in the transduction assay to reduce the chance of transformation.
Titering of the transducing lysate Protocol
1. Prepare a culture of the indicator strain in nutrient medium and grow to midlog phase. 2. Make a serial dilution of the phage lysate in 0.5 M rris buffer just before use.
3. Mix 100 ~Jl of each phage dilution with 1 ml of indicator bacteria in soft agar and pour immediately onto an agar plate as described for production of phage particles. 4. Enumerate plaque forming units after overnight incubation.
Transduction
Protocol
1. Mix 10-100 ml of log-phase recipient cell culture with transducing phage particles at MOIs ranging from 0.01 to 5. 2. Incubate the mixture at room temperature for 10 min to allow phage adsorption. 3. Remove the unabsorbed phages by three rounds of centrifugation and washes with artificial seawater. 4. Resuspend the final cell pellet in 0.5-1.5 ml of nutrient broth, and allow the cells to recover in this nonselective medium for 10-20 min before plating onto nutrient plates selective for the genetic determinants serving as markers for transduction. 5. The transducing lysate (containing no recipient) and recipient only should also be plated onto the same selective plates as controls. 6. Incubate the plates for two to six days before counting colonies of transductants. Longer incubation periods may be needed for slowgrowing marine bacteria. Extended incubation is often necessary to allow for the phenotypic expression of the transduced gene. 7. The frequency of transduction can be expressed as transductants per transducing phage or per recipient. 120
Transduction in natural populations Natural marine bacterial populations can be used directly as recipients for transduction if proper genetic markers (i.e. an unique plasmid) are used for the donor bacteria (Figure 7.3). The production of transducing lysates should be the same as for transduction in the cultured system described above. Compared to recipients in culture, bacteria in natural seawater are much less abundant. Therefore, the bacterial community should be concentrated to allow detection of transduction.
Equipment and supplies •
Membrex Rotary Biofiltration Device, equipped with 100 KD filter, or other cell concentrating device (tangential flow, etc.) • All materials and supplies for transduction assay in the cultured bacteriophage and host systems
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Protocol
1.
2.
3. 4. 5.
6.
7.
Concentrate 20 to 100 1 of seawater from offshore environments using a Membrex vortex flow filtration system (Jiang et al., 1992) to 50 ml. Mix 10 ml of this concentrate with various concentrations of transducing particles and allow phage to adsorb onto the bacteria at room temperature for 10 min. Filter the mixture onto a 0.2 ~m filter and rinse with sterile artificial seawater to wash off the unadsorbed phages. Resuspend cells collected on the filter in 2 ml of nutrient broth and plate 100 ~tl on selective medium plates. Plate an equal volume of concentrated sample without addition of transducing phages and transducing lysate alone as negative controls. Incubate for at least 4 days before enumeration of transductants. Many indigenous marine bacteria are resistant to various antibiotics. If an antibiotic resistant plasmid is used as a genetic marker for transduction, colony hybridization with a plasmid specific probe can be performed to distinguish the transductants from indigenous resistant bacteria. The frequency of transduction can be expressed as transductants per transducing phage particle.
FUTURE DIRECTIONS In terms of lysogeny, the factors which control the regulation of this phenomenon in the environment will hopefully be determined by future research. That is, what environmental conditions control the lysogenic decision upon infection with a temperate phage? Does pseudolysogeny 121
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play a role in production of phage in the marine environment? And, do p h y t o p l a n k t o n blooms crash because of induction of temperate algal viruses? Some of these questions are experimentally difficult to answer with current technology, while others have yet to be investigated, In comparison with freshwater environments, m u c h less effort has been directed at the investigation of transduction in marine waters. Many methods that were designed for freshwater habitats are also suitable for the investigation of in situ transduction in marine environments. Examples of these methods include: (1) transduction assays in flow through environmental chambers that are incubated at ambient temperature; (2) transduction with spontaneous induced phage without separation and purification from d o n o r bacteria (i.e. mixing the d o n o r and recipient in an environmental chamber). In addition, several new approaches that m a y extend our current understanding of transduction in the marine environment are also w o r t h y of investigation. First, the native marine bacteriophages can now be easily concentrated and purified from seawater. It should be interesting to investigate the frequency of transduction by these indigenous marine bacteriophages. In this transduction system, auxotrophic bacteria can be used as recipients as per Chiura (1997). Secondly, all transduction assays to date have been designed to detect the gene transfer event in cultivable marine bacteria. Since less than 1% of marine bacteria are culturable by current methods, it is important to develop strategies to detect transduction events in non-cultivable marine bacteria. One of the strategies is to use a fluorescent in situ hybridization technique (FISH) to trace the uptake of genetic markers in a single cell without cultivation. Alternatively, in situ PCR can also be used to increase the detection sensitivity. Additionally, plasmids containing the green fluorescent protein (GFP) gene can be used, and transduction detected by epifluorescence microscopy (Dahlberg et al., 1998). In conclusion, marine transduction is still a y o u n g and growing field of research. As new techniques are developed to study gene transfer in natural populations, the overall importance of this process in the evolution of microbial populations in the environment will unfold.
References Ackermann, H. W. and DuBow, M. S. (1987). Viruses qf Prokaryotes. Vol. 1. General properties of bacteriophages. CRC Press, Boca Raton, FL. Amin, M. K. and Day, M. J. (1988). Donor and recipient effects on transduction frequency in situ. REGEM1 Program. Baross, J. A., Liston, J. and Morita, R. Y. (1978). incidence of Vibrio parahaemolyticus bacteriophages and other Vibrio bacteriophages in marine samples. Appl. Environ. Microbiol. 36, 492-499. Benzinger, R. and Hartman, P. E. (1962). Effect of ultraviolet light on transducing phage P22. Virology 18, 614-626. Borsheim, K. Y., Gratbak, G. and Heldal, M. (1990). Enumeration and biomass estimation of planktonic bacteria and viruses by transmission electron microscopy. Appl. Environ. Microbiol. 56, 352-356.
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Buck, G. A. and Groman, N. B. (1981). Genetic elements novel for Corylwbacterium diphtheriae: specialized transducing elements and transposons. I. Bacteriol. 148, 143-152. Cavenagh, M. M. and Miller, R. V. (1985). Specialized transduction of PseudomoHas aeruy,il~osa PAO by bacteriophage D3. J. Bacteriol. 165, 448-452. Chiura, H. X. (1997). Generalized gene transfer by virus-like particles from marine bacteria. Aquat. Microb. Ecol. 13, 75-83. Chirua, H. X., Hato, K., Hiraishi, A. and Maki, Y. (1998). Gene transfer mediated by virus of novel thermophilic bacteria in hot spring sulfur-turf microbial mats. Eighth International Syrnposium on Microbial Ecology (1SME-8). Program and Abstracts. Cochran, P. K. and Paul, ]. H. (1998) Seasonal abundance of lysogenic bacteria in a subtropical estuary. Appl. EH~qrolL Microbi~d. 64, 2308-2312. Cochran, P. K., Kellogg, C. A. and Paul, J. H. (1998) Prophage induction of indigenous marine lysogenic bacteria by environmental pollutants. Mar. Ecol. Progr. Set. 164, 125-133. Dahlberg, C., Bergstrom, M. and Hermansson, M. (1998). hz situ detection of high levels of horizontal plasrnid transfer in marine bacterial communities. Appl. D~viron. Microbiol. 64, 2670-2675. Denhardt, D. T., Dressier, H. H. and Ray, D. S. (eds). (1978). Sirrah' stra~ded DNA Pha~es. Cold Spring Harbor Laboratory. Dr6ge, M., Pi_ihler, A. and Selbitschka, W. (1998). Horizontal gene transfer as a biosafety issue: a natural phenomenon of public concern. J. Biotech. 64, 75-90. Germida, J. J. and Khachatourian, G. G. (1988). Transduction of Escherichia colt in soil. CaJl. J. Microbiol. 34, 190-193. Jain, R., Rivera, M. C. and Lake, J. A. (1999). Horizontal gene transfer among genomes: the complexity hypothesis. Proc. Natl. Acad. Sci. USA 96, 3801-3806. Jarolmen, H., Bonke, A. and Crowell, R. I. (1965). Transduction of Staphylococcus aim'us to tetracycline resistance iH vivo. J. Bacteriol. 89, 1286-1290. Jensen, E. C., Schrader, H. S., Rieland, B., Thompson, T. L., Lee, K. W., Nickerson, K. W. and Kokjohn, T. A. (1998). Prevalence of broad-host-range lyric bacteriophages of Sphaerotilus izatalls, Escherichi colt, and Pseudomonas aeruliJwsa. Appl. DzviroH. Microbiol. 64, 575-580. ]tang, S. C. and Paul, J. H. (1994). Seasonal and diel abundance of viruses and occurrence of lysogeny/bacteriocinogeny in the marine environment. Mar. Ecol. Progr. Ser. 104, 163 172. Jiang, S. C. and Paul, ]. H. (1996). Occurrence of lysogenic bacteria in marine microbial communities as determined by prophage induction. Mar. Ec~d. Prog. Set. 142, 27-38. ]tang, S. C. and Paul, J. H. (1998a). Significance of lysogeny in the marine environment: studies with isolates and a model for viral production. Microb. Ecol. 35, 235-243. Jiang, S. C. and Paul, J. H. (1998b). Gene transfer by transduction in the marine environment. Appl. E1zvirou. Microbiol. 64, 2780-2787. ]tang, S. C., Thurmond, J. M., Pichard, S. L. and Paul, ]. H. (1992). Concentration of microbial populations from aquatic environments by vortex flow filtration. Mar. Ecol. ProS. Set. 80, 101 107. Keynan, A., Nealson, K., Sideropoulos, H. and Hastings, J. W. (1974). Marine transducing bacteriophage attacking a luminous bacterium. J. Wr01. 14, 333340. Kidambi, S. P., Ripp, S. and Miller, R. V. (1993). Evidence for phage-mediated transfer among Pseluh~moHas aCI'HgiHosI7 o n the phylloplane. Appl. D~virotl. Microbiol. 60, 496-500.
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Kokjohn, T. A. (1989). Transduction: mechanism and potential for gene transfer in the environment. In: Gene Transfer in the Environment (S. B. Levy and R. V. Miller, Eds), pp. 73-97. McGraw-Hill, New York. Levin, B. R. and Lenski, R. E. (1983). Coevolution in bacteria and their viruses and plasmids. In: Coevolution (D. J. Futuyma and M. Slaktin, Eds), pp. 99-127. Sinauer Associates, Inc., Sunderland, MA. Levisohn, R., Moreland, J. and Nealson, K. H. (1987) Isolation and characterization of a generalized transducing phage for the marine luminous bacterium Vibrio fischeri MJ-I. 1. Gen. Microbiol. 13, 1577 1582. Miller, R. V. (1992). Methods for evaluating transduction: An overview with environmental considerations. In: Microbial Ecolo\~y: Principles, Methods, ~Tud Applicatiopls (M. A. Levin, R. J. Seidler and M. Rogul, Eds), pp. 229-251. McGraw-Hill, New York. Miller, R. V., Ripp, S., Relicon, J., Ogunseitan, O. A. and Kokjohn, T. A. (1992). Virus-mediated gene transfer in freshwater environment. In: Gene Transfers nnd Etlviromnent (M. J. Gauthier, Ed.), pp. 51-62. Springer-Verlag, Berlin. Morgan, A. E (1979). Transduction of Pseudomonas aeruginosa in a freshwater environment. Appl. Environ. Microbiol. 36, 724-730. Morrison, W. D., Miller, R. V. and Sayler, G. S. (1978). Frequency of Fl16 mediated transduction of Pseudomonas aerugiuosa in a freshwater environment. Appl. EHviron. Microbio[. 36, 724-730. Novick, R. P. and Morse, S. I. (1967). hi vivo transmission of drug resistance factors between strains of Staphyh~coccus aureus. J. Exp. Med. 125, 45-49. Novick, R. P., Edelman, I. and Lofdahl, S. (1986). Small Staphylococcus aureus plasmids are transduced as linear multimers that are formed and resolved by replicative process. J. Mol. Biol. 192, 209-220. Paul, J. H. and Myers, B. (1982). The fluorometric determination of DNA in aquatic microorganisms employing Hoechst 33258. Appl. Ewe,iron. Microbiol. 43, 1393-1399. Ripp, S. and Miller, R. V. (1995) Effects of suspended particulates on the frequency of transduction among Pseudomonas neruginosa in a freshwater environment. Appl. D~viroH. Microbio[. 61, 1214-1219. Ripp, S., Ogunseitan, O. A. and Miller, R. V. (1994). Transduction of a freshwater microbial community by a new Pseudomonas aeruy,inosa generalized transducing phage, UT1. Mol. Ecol. 3, 121-126. Saye, D. J. and Miller, R. V. (1989). The aquatic environment: consideration of horizontal gene transmission in a diversified habitat. In: Gene Transfer ii~ the Ei1vironmel~t (S. B. Levy and R. V. Miller, Eds), pp. 223-259. McGraw-Hill, New York. Saye, D. J., Ogunseitan, O., Sayler, G. S. and Miller, R. V. (1987) Potential for transduction of plasmids in a natural freshwater environment: effect of donor concentration and a natural microbial community on transduction in PseudomoJlas aeruginosa. Appl. Environ. Microbiol. 53, 987-995. Saye, D. J., Ogunseitan, O. A., Sayler, G. S. and Miller, R. V. (1990). Transduction of linked chromosomal genes between Pseudomonas aeruginosa during incubation in situ in a freshwater habitat. Appl. Environ. Microbiol. 56, 140-145. Scanlan, D. J. and Wilson, W. H. (1999). Application of molecular techniques to addressing the role of P as a key effector in marine ecosystems. Hydrobiol. 401, 149-175.
Schichklmaier, P. and Schmieger, H. (1995). Frequency of generalized transducing phages in natural isolates Salmonella typhimurium complex. Appl. Environ. Microbiol. 61, 1637-1640. Sternberg, N. L. and Maurer, R. (1991). Bacteriophage-mediated generalized trans-
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duction in Escherichia coli and Sahnonella typhimurium. Method Enzymol. 204,
19-43. Stozky, G., (1989). Gene transfer among bacteria in soil. In: Ge~w Traus~er in the ElzviromHent (S. B. Levy and R. V. Miller, Eds), pp. 165-222. McGraw-Hill, New York. Tapper, M. A. and Hicks, R. E. (1998) Morphology and abundance of free and temperate viruses in Lake Superior. Limnol. Occauo?¢r. 43, 95-103. Waterbury, J. B. and Valois, E W. (1993). Resistance to co-occurring phages enables marine Synechococcus communities to coexist with cyanophages abundant in seawater. Appl. EuviroJ~. Microbiol. 59, 3393-3399. Weinbauer, M. G. and Suttle, C. A. (1996). Potential significance of lysogeny to bacteriophage production and bacterial mortality in coastal waters of the Gulf of Mexico. Appl. E1tviroJt. Microbiol. 62, 4374-4380. Weinbauer, M. G. and Suttle, C. A. (1999). Lysogeny and prophage induction in coastal and offshore bacterial communities. Aquatic Microbial Ecol. 18, 217-225. Wichels, A., Biel, S. S., Gelderblam, H. R., Brinkhoff, T., Muyzer, G. and Sh(itt, C. (1998). Bacteriophage diversity in the North Sea. Appl. E~viron. Microbiol. 64, 4128-4133. Wilcox, R. M. and Fuhrman, J. A. (1994). Bacterial viruses in coastal seawater: lytic rather than lysogenic production. Mar. Ecol. Prog. Ser. 114, 35M5. Wilson, W. H., Turner, S. and Mann, N. H. (1998) Population dynamics of phytoplankton and viruses in phosphate-limited mesocosm and their effect on DMSP and DMS production. Estuar. Coastal Shelf Sci. 46, 49-59. Wommack, K. E. and Colwell, R. R. (2000). Viroplankton: viruses in aquatic ecosystems. Microbiol. Mol. Biol. Rev. 64, 69-114. Zeph, L. R. and Stotzky, G. (1989). Use of a biotinylated DNA probe to detect bacteria transduced by bacteriophage P1 in soil. Appl. E1~viro11. Microbiol. 55, 661-665. Zeph, L. R., Onaga, M. A. and Stotzky, G. (1988). Transduction of Escherichia coil by bacteriophage P1 in soil. Appl. Environ. Microbiol. 54, 1731-1737. Zinder, N. D. and Lederberg, J. (1952). Genetic exchange in Salmoi~ella. ]. Bactcriol. 64, 679-699.
List of suppliers Fisher Scientific
Sigma Chemical Corporation
3970 Johns Ctvek Court Suwauee, GA 30024, USA 1-800-766-7000
PO Box 14508 St. Louis, M O 63178, USA 1-800-325-3010
Anodisc and Nuclepore filters, microtiter plates
Electron microscopy grade glutaraldehyde Mitomycin C
Micron Separations Inc. 135 Flanders Road PO Box 1046 Westborou~h, M A 01581, USA 1-800-444-8212 M e m b r e x / O s m o n i c s Rotary Biofiltration Device
Difco Laboratories P. 0. Box 331058 Detroit, MI, 48232-7058, USA Phone: 800-521-0851 Fax: 313-462-8517 Microbiological media
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8 Enumeration of Total and Highly Active Bacteria Barry Sherr', Evelyn Sherr' and Paul del Giorgio 2 'College of Oceanic and Atmospheric Sciences, 104 Ocean.Admin. Bldg, Corvallis, OR 97331-5503, USA ~Horn Point Laboratory, Univ. of Maryland Center for Environmental Studies, P.O.Box 775, Cambridge, MD 21613, USA
CONTENTS Introduction Methods Results Discussion and future directions
~I,~,~,~,~,~I, I N T R O D U C T I O N Counting bacteria in natural environments has been a long-standing endeavor for aquatic microbial ecologists. Abundance and biomass of bacteria are central parameters to understanding the roles of heterotrophic microbes in marine ecosystems. Assessment of bacterial abundance in seawater has evolved through several stages: (1) Enumeration of culturable bacteria based on ability of single bacterial cells to form colonies on marine agar plates. (2) Enumeration of total bacteria based on universal fluorochrome staining of cells and epifluorescence microscopy. (3) Enumeration of metabolic and phylogenetic categories of bacterial cells, based on use of specifically targeted fluorochromes and molecular probes, via epifluorescence microscopy and flow cytometry. Besides direct counting methods, bulk chemistry approaches have also been used for quantifying prokaryotic biomass; these include analysis of phospholipids specific to bacteria (White et al., 1979) and the Limulus amoebocyte lysate assay for analysis of lipopolysaccharide, a constituent of the cell walls of Gram-negative bacteria (Watson et al., 1977; Karl and Dobbs, 1998). Early approaches to enumeration of bacteria in the sea included direct counts with transmitted light microscopy and estimates of the number of culturable bacteria by plating methods (Wood, 1965). In plating assays, the water sample is appropriately diluted, and the number of colonies METHODS IN MICROBIOLOGY, VOI,UME 30 ISBN 0-12-521530-4
(_opyright © 2001 Academic Press Ltd All rights of reproduction in any form rcserx vd
(which in theory arise from individual bacterial cells) that appear after a 3-5 day incubation on solid agar plates are counted. Transmitted light microscopy methods were not very satisfactory; and the plate count method yielded low estimates of the number of bacteria in seawater, on the order of 10-"to 10~ ml ~. The discrepancy between the low abundance based on plate counts of 'culturable' or 'viable' bacteria, and the I0 to 100fold higher abundance of bacterial cells that can be counted directly in seawater has resulted in controversies that are still largely unresolved. A revolution in marine microbiology occurred with the development of direct count methods using fluorescent stains and epifluorescence microscopy. Strugger (1948) used the fluorochrome acridine orange (AO) to visualize soil bacteria. Methods for routine inspection of fluorochromestained bacteria were delayed until the introduction of membrane filters for collection of bacteria, combined with epifluorescence microscopy (Zimmermann and Meyer-Reil, 1974). Hobbie et al. (1977) proposed filtration of seawater bacteria onto irgalan black-stained 0.2 bun pore-size membrane filters, and staining the filtered cells with AO. This method was widely adopted, and has since yielded abundant information on the distribution of bacterial cells in natural waters (Bird and Kalff, 1984; Newell et al., 1986; Cole et al., 1988). Porter and Feig (1980) subsequently suggested use of the DNA stain 4'6-diamidino-2-phenylidole (DAPI) for visualizing bacterial cells, the advantage being low background (nonspecific) fluorescence compared to AO. These two stains are still commonly used to enumerate bacterial cell abundance in the sea. Bacterial enumeration entered a new phase with the application of flow cytometry, the availability of an array of specifically targeted fluorochromes, and development of taxon-specific molecular probes. It is now possible to quantify classes of cells with specific physiological and phylogenetic characteristics. In this chapter we will cover standard methods of enumeration of total bacterial numbers with epifluorescence microscopy, discuss the advantages of flow cytometery over microscopic enumeration, present conversion factors used to estimate carbon biomass based on cell abundance and cell size, and also describe the CTC assay which identifies bacteria with highly active electron transport systems (ETS). Other methods to detect cell-specific physiological status of bacterial cells are presented in Howard-Jones et al. (Chapter 10, this volume).
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METHODS
Handling of samples for bacterial enumeration Samples containing bacteria should be preserved before filtration. Fixation with a final volume of 1% to 6% formalin is commonly used. We normally preserve samples with 2% final volume borate-buffered formalin (20 btl ml -~of sample); however Choi et al. (1996) recommended 10% final volume of formalin (100 gl ml ' of sample) and letting samples stand for 10 minutes at room temperature when bacterial cells will be
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subjected to harsh treatment, e.g. washing with warm propanol to detect cells with or without visible nucleoids. Turley (1993) alternatively suggested preserving samples with 2.5~7~ final concentration glutaraldehyde using 0.2-btm-filtered 25% SEM grade glutaraldehyde (100 btl ml ' of sample). The number of bacteria declines with time in aldehyde-fixed liquid samples (Turley, 1993). Gundersen et al. (1996) suggested that one reason is enzymatic lysis of cells, even in preserved samples. Vosjan and van Noort (1998) found that most of the bacteria that disappeared during storage of liquid samples were cells that did not have visible nucleoids; abundance of bacteria with visible nucleoids was constant over a 70 day storage period. For microscopic preparations, samples should be processed within a day, and mounted filters either immediately inspected or kept frozen (-20°C) in slide boxes until counts are made. For the CTC assay preserved liquid subsamples (1-2 ml) should be quick-frozen in liquid nitrogen, and stored frozen at -20°C (the quick-freeze step is not absolutely critical). For flow cytometry, 1 ml samples are fixed with 1-2% final concentration of glutaraldehyde for 10 to 60 minutes, then stored frozen until thawed and immediately analyzed (Olson et al., 1993; see Flow cytometry section). Freezing samples and storing samples for flow cytometry in liquid nitrogen is recommended (Olson et al., 1993).
Epifluorescence microscopy Materials required
Equipment •
Microscope outfitted for epifluorescence microscopy, with appropriate filter sets for the fluorochromes of interest, a 50 W or 100 W mercury lamp or 100 W zenon lamp, a 100× oil immersion objective lens, and an ocular grid (graticule) of I 0 × I 0 squares • Filtration apparatus for 25 mm filters • Millipore forceps (In our experience, Millipore forceps, which have broad blunt edges, are superior for manipulation of membrane filters) • Adjustable pipettors for quantitative delivery of water samples and fluorochromes • Stage micrometer for measuring width of microscope fields of view • -20°C freezer for storage of prepared slides
Supplies •
Black-stained 0.2~m pore-size 25 mm polycarbonate membrane filters, Nuclepore or PoretJcs • Cellulosic backing filters to support the membrane filters and aid in even dispersion of cells over the membrane filter surface. Commonly used backing filters are 0.45 ~m MJllJpore filters or 0.8 ~m Nuclepore Membra-fil filters • Gelman Supor TM AcrodJscs TM, 0.2 ~Jm, or equivalent • Glass slides and 25 mm square No. I cover slips
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• • • • •
Immersion oil, e.g. Cargille type A or type LF; or Resolve TM low viscoscity immersion oil Pens that can mark on plain or fritted glass slides Slide boxes, preferably with metal snap closures 20 ml disposable plastic syringes 1.5 ml plastic vials, e.g. microfuge tubes or cryovials
Solutions
Note: precautions, e.g. wearing gloves, should be taken w h e n handling DNA and RNA-binding fluorochromes; these c o m p o u n d s are potential carcinogens. • Acridine orange stock solution, 0. I% AO, 6% formalin, 0.2 lum filtered. Dissolve 20 mg of AO in 200 ml of 0.2-lum-filtered seawater or artificial seawater of the approximate salinity of the samples, shake or stir to completely dissolve. Filter the AO solution through a 0.2 pm filter (note AO stains glassware, so it is a good idea to reserve a glass filtration unit: flask, bottom, and tower, specifically for this use). Pour the AO solution into a dark 250 ml glass bottle and add 12 ml of 37% formaldehyde (formalin). Stored in a refrigerator, the AO stock solution is good for months.
When processing samples, it is useful to d r a w up -15-18 ml of AO stock solution into a 20 ml plastic syringe and attach a 0.2 pm Gelman Acrodisc to the tip of of the syringe. Then, freshly 0.2-pm-refiltered AO solution can be neatly dispensed onto the filters as needed. This works especially well for shipboard sample preparation. •
DAPI stock solution, 200 lug ml ' (0.57 mM). Dissolve I 0 mg of DAPI in 50 ml of distilled water, filter through a 0.2lure Gelman Acrodisc, using a plastic syringe. It is convenient to make batches as DAPI can be bought in 10 mg lots, and there is no need for weighing milligram quantities with this approach. Dispense I-2 ml aliquots of the DAPI stock into labeled plastic vials, e.g. 1.5-2 ml microcentrifuge tubes or cryovials, store in a -20°C (or lower temperature) freezer. DAPI stock is good for up to a year in the freezer. • Borate-buffered formalin. Fill a 500 ml dark glass bottle with about 450 ml of formalin (37% formaldehyde, reagent grade). Add sodium borate crystals until an undissolved layer of crystals lies on the bottom, i.e. a saturated solution. When preserving samples, the formalin should be freshly filtered through 0.2 pm.We dispense formalin into samples using a plastic syringe fitted with a 0.2 lum Acrodisc. Procedures Acridine orange direct counts (AODC)
Sample preparation 1. Prepare a glass slide. Label one end of the slide with the sample code: e.g. name or number. Put a small drop of immersion oil onto the slide and smear fiat with the edge of a cover slip or with a pipette tip. When we prepare duplicate subsamples, we normally m o u n t both filters,
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4.
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6.
7.
with two separate cover slips, on one glass slide to conserve storage space in slide boxes. Place a 0.8 bun or 0.45 btm cellulosic backing filter onto a 25 m m filtration bottom. Wet with a few drops of deionized water. (Note: the same backing filter can be used for multiple samples; replace daily or if torn.) Place a black 0.2 btm m e m b r a n e filter, shiny side up, onto the wetted backing filter. Prerinse the filtration tower with 0.2-btm-filtered seawater, shake dry, and carefully place on top of the filter. Clamp tightly. If the tower is not centered and clamped properly, the sample m a y leak out. Filter an aliquot of preserved sample through the 0.2 ~1nl black m e m b r a n e filter. Sample v o l u m e d e p e n d s on bacterial concentration. For oligotrophic samples in which bacterial abundance is less than 5 x 10 ~ ml ', 5-10 ml per sample is necessary. For eutrophic samples in which bacterial a b u n d a n c e is greater than 10"ml ', 1-2 ml per sample is adequate. For bacterial cultures with abundances in excess of 10: cells ml ~, it m a y be necessary to dilute samples with 0.2-btm-filtered seawater. The v o l u m e filtered should be at least 1 ml to ensure even distribution on the filter. Release the v a c u u m and gently add 1 ml of 0.2-btm-filtered AO solution onto the top of the filter. Run the AO solution d o w n the side of the filter tower to minimize disturbance of the cells on the filter. It is important to have as uniform distribution of cells as possible for accurate counts. Allow to stand for 2 rain, and then turn on the v a c u u m and filter d o w n the AO solution. An alternative m e t h o d r e c o m m e n d e d by Turley (1993) inw)lves filtering a sample d o w n to 2 ml, adding 200 btl of a 1% AO stock solution (10-told more concentrated than the AO stock used here), and letting it stand for 5 min before filtration. This approach avoids the possibility of artefactual redistribution of cells on the filter when the AO solution is added. With the v a c u u m still on, carefully remove the m e m b r a n e filter from the u n d e r l y i n g backing filter. If the v a c u u m is released, fluid will flow back u p through the pores of the filter, dislodging some of the cells on the surface. Lay the filter, sample side up, onto the film of immersion oil on the slide. Put one drop of immersion oil onto the center of the filter, and gently lay a cover slip on top of the filter, being careful to avoid air bubbles. For each set of filters, prepare one or more blank filters by filtering d o w n 1 ml of AO stock solution without a sample. These will be used to obtain a value for background counts. Inspect the sample immediately, or place the slide in a slide box in a 20°C (or lower temperature) freezer. The samples are good for several months at least. We have counted samples up to a year old. We have also lost samples due to freezer malfunction. It is best to do the counts as soon as possible.
Bacterial e n u m e r a t i o n
1. Obtain a factor for the n u m b e r of microscope grids per filter for the particular filter apparatus and microscope being used. Measure the 133
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diameter in millimeters of the bottom of the filter tower, and calculate the area in square millimeters (ram2). This is the area of the sample on the surface of the filter. Next, determine the width of each side of the 10 x 10 ocular grid. Put a drop of immersion oil on the ram-long scale of a stage micrometer. Focus on the micrometer scale with the lOOx lens. Width between lines of the micrometer is 10 ~m. Line up one side of the ocular grid with the micrometer scale and determine the length in microns. Convert this length to millimeters (1000 microns per mm), and square this value to obtain the area in square millimeters (mm ~) of the grid field. Divide the sample area by the grid area to arrive at the factor for n u m b e r of grids per filter (usually on the order of several × 10~). 2. Use a blue light epifluorescence filter set to visualize AO-stained bacteria, e.g. Zeiss filter set 47 77 09 (BP 450-490 excitation filter, FT 510 beam splitter and LP 520 barrier filter). Find the surface plane of the filter that contains the stained bacteria. This step is not always easy. One trick is to illuminate the filter with a low level of transmitted (white) light, focus on the m e m b r a n e pores at 100x, and then turn off the transmitted light and find the plane containing the bacteria with epifluorescence illumination using the fine focus adjustment. If the microscope is equipped with parfocal lenses, one can also focus on the surface of the filter using a lower magnification lens, and then switch to the 100x lens. 3. For each sample, count the n u m b e r of bacteria in a whole grid for 7 to 10 r a n d o m l y chosen fields distributed over the filter. An appropriate density of bacteria on the filter surface would result in about 30 to 50 bacterial cells within a 10 x 10 ocular grid at 1000x magnification (100x objective lens and 10x ocular magnification). Ideally, a m i n i m u m of 300 bacteria will be counted per filter (Kirchman, 1993); Turley (1993) suggested counting up to 600 cells per filter (14 to 20 fields). Kirchman et al. (1982) and Kirchman (1993) r e c o m m e n d counting 7 to 10 fields on two replicate filters for increased precision; this is the approach that we take in our research. The counts from the two filters are combined to calculate a single abundance value. Typically about 1-2 x 10" bacteria will be on the surface of a filter. The 300 to 600 cells counted per filter thus represents only 0.03% to 0.06% of the total n u m b e r of bacteria. This is the basis of concerns regarding uniform distribution of cells on the filter, as well as of the large coefficients of variation for estimates of bacterial abundance obtained via microscopic counts (Kirchman, 1993). 4. Determine the average n u m b e r of bacterial cells per grid for each sample counted. Also count cells on the filters prepared without a d d e d sample. Calculate bacterial abundance from the equation: cells ml ' = [(sample cells per grid - background cells per grid) x grids per filter] / v o l u m e of sample. In order to determine an average cell abundance, several replicate subsamples from the same water sample should be counted.
134
DAPI direct counts
Using DAPI rather than AO for bacterial enumeration has the advantages of low background fluorescence, less interference from photopigments, and the fact that DAPI fluorescence does not fade while a field is being counted. While AO stains both DNA and RNA, DAPI supposedly stains only DNA. However, Zweifel and Hagstrom (1995) demonstrated that for aldehyde-fixed marine samples, DAPI did not stain DNA very efficiently, but did stain other components of bacterial cells non-selectively. Staining bacteria with DAPI at lower salinity, and removing DAPI not bound to DNA via a propanol rinse, allowed detection of the nucleoid region in bacterial cells (Zweifel and Hagstrom, 1995; see description of the visible nucleoid method in Chapter 10 of this volume). Direct counts using DAPI require a UV light excitation filter set, e.g. Zeiss filter set 47 77 02 (G 365 excitation filter, FT 395 beam splitter and LP 420 barrier filter). It is also important that the objective lens is Neofluor, i.e. does not have coatings that preclude transmission of UV light. The procedures are generally the same as for the AODC method. Use a separate filter apparatus for DAPI, otherwise there may be AO contamination of the filter. Steps 3 and 4 of the sample preparation protocol outlined above are modified for DAPI counting as follows: 3. Thaw a 1-2ml vial of DAPI solution. Filter sample down to 2ml volume, release vacuum. 4. Add 50 btl of DAPI solution to the sample to yield a final concentration of 5 btg DAPI ml ' (14 btM)sample, cover with aluminum foil to keep out light, and allow to stand for 7 minutes. Filter down and process as for the AO filters. Note: final concentrations of DAPI of 1-10 btg m l ' (3-29 [.IM) of sample are usually recommended (Velji and Albright, 1993; Hoff, 1993; Kawai et al., 1999). The DAPI concentration of 0.01 btg mU ( 0.03 btM) originally suggested by Porter and Feig (1980) is much too low and leads to underestimates of cell number (Hoff, 1993; Kawai et al., 1999). Subsamples can be pre-stained with DAPI in a vial in advance of filtration; after 7 minutes, the samples remain optimally stained for hours. Sizing bacterial cells
Empirical studies have shown that the carbon content of marine bacteria varies with size (Norland, 1993; Table 1). Determination of the average cell volume for bacteria in each habitat or water mass sampled is recommended in order to estimate carbon biomass of bacteria. Several approaches have been used (Bratbak, 1993). The simplest is photomicrography, in which photographs are taken of bacterial cells in a number of fields (5-10) on a filter using fine-grained black and white film (Fuhrman, 1981; Bratbak, 1993; Lee, 1993). A photograph of the scale of a stage micrometer at 1000x is also made. After the film is developed, the negatives are mounted in frames and projected onto a flat surface. A ruler is calibrated using the photograph of the stage micrometer line spacing (10 ~tm). Individual bacterial cells that are in sharp focus are then 135
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measured with the ruler, and length and width recorded for each. Edge discrimination, which is a factor in image analysis of bacterial cells, is generally not a problem with the photographic method. Cell biovolume is calculated using the formula for a prolate spheroid: V = 0r/4)W ~(L - W/3), where W = cell width and L = cell length. A sufficient number of cells are measured to yield a reasonably low standard error of the mean value. Once the average cell size has been determined, carbon biomass can be calculated using an appropriate factor based on empirical determinations of bacterial carbon content (Norland, 1993; Table 1). Other approaches to cell sizing involve electron microscopy (Bratbak, 1993), imaging cytometry (Sieracki et al., 1989; Sieracki and Viles, 1998), X-ray microanalysis (Heldal, 1993), Coulter Counter analysis (Robertson et al., 1998), and flow cytometry (see next section). While a variety of approaches has yielded similar estimates of cellspecific carbon contents (-10-30fgC per cell), cell biovolume-specific factors appear to be influenced by the method used to size bacterial cells. Microscopy-based biovolumes of in situ marine bacteria are in the range 0.03-0.15 gm ~(Table 8.1, Viles and Sieracki, 1992; Suzuki et al., 1993; Sherr et al., 1997). Choice of fluorochrome can influence size estimates. Suzuki et al. (1993) reported that cell volumes obtained from DAPI-stained bacteria were on average only 59c7~ of cell volumes determined from AOstained bacteria. Other approaches, such as X-ray microanalysis and particle counting, yield cell biovolumes about 3-fold higher, 0.10-0.30 pm -~,compared to microscopy-based biovolumes (Table 8.1). Based on current knowledge, the factors of 20 fg C per cell suggested by Lee and Fuhrman (1987), or of 12 fg C per cell for bacterioplankton in oligotrophic waters and 30 fg C per cell for bacterioplankton in eutrophic waters suggested by Fukuda et al. (1998) are appropriate for studies in which average cell size was not determined. For cases in which bacterioplankton cell size has been determined by microscopic analysis, the biovolume-specific conversion factors of Simon and Azam (1989) can be used.
Flow c y t o m e t r y General principles Basis of method During the last decade, flow cytometry has become a standard approach to enumeration of heterotrophic and autotrophic bacteria in aquatic systems (Chisholm et al., 1988; Olson et al., 1991, 1993; Monger and Landry, 1993; Button and Robertson, 1993; Li et al., 1995; Davey and Kell, 1996; Binder and Liu, 1998). With flow cytometry, a liquid sample containing the target cells (i.e. bacterial or algal cells) is entrained within the sheath flow and the particles are aligned so that they intercept a focused beam of light, usually a laser, in single file. Upon illumination with the laser, each particle scatters light at different angles, and
136
Table 8. I Estimates of carbon content: fg (10 '~g) C per cell, and pg (I 0,2 g) C gm ~ of marine bacteria fg C per cell
pgC ~lm3
Reference
0.036-0.073
20 + 0.8
0.35
Lee and F u h r m a n 1987
Southern California, USA coastal water (microscopy + carbon content of macromolecules)
0.026 0.036 0.050 0.070 0.100 0.200 0.400
10.4 12.6 15.2 18.7 23.3 35.0 53.5
{).40 0.35 0.30 0.27 0.23 0.17 0.13
Simon and A z a m 1989, Table 5
N o r w e g i a n coastal w a t e r (X-ray microanalysis)
{).15-0.6
15-42
0.06-0.13
Tuomi et al. 1995 Table 1
Habitat
(analytical approach) Long Island, USA beach surf
Cell volume
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S c a n d i n a v i a n coastal waters (X-ray microanalysis)
0.11 0.20-0.21 0.28 {).31
9.0 7-12 19 3l
0.08 {).03-0.06 0.07 0.10
Fagerbakke et al. 1996, Table 1 ¢o
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I.
{} Central Arctic Ocean, 0-100 m (microscopy + conversion factors from Table 5 of S i m o n and A z a m 1989)
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Oceanic bacterial isolates: {i. 14C acetate uptake)
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0.29_+0.02
10.3±1.8
Sherr eta/. 1997, Table 3
Robertson et aI. 1998, Table 4
Cyclochzsticus oligotrophus Coulter C o u n t e r + flow cytometry)
Cycloclasticus oligotrophus
0.13 0.16 0.24-0.25
13.3" 16.8 19.4-27.1
0.10 0.10 0.08-0.11
Marinobach'r sp strain T2
0.22-{l.24
21.7-25.4
0.10
(ii.
O p e n ocean vs. coastal (HTCO** analysis) Oceanic sites Coastal sites
12.4_+6.3 30.2 + 12.3
*Calculated as 47!~ of dry weight cell ' **HTCO= high temperature catalytic oxidation
137
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may also emit fluorescence. Light scattered and emitted from each particle is captured by a set of photomultiplier tubes or photodiodes that convert the photon flux into an electrical signal which can then be digitized and analyzed with appropriate software. In this way, individual particles, most often cells, cannot only be detected and enumerated, but the optical parameters analyzed can be used to assess a host of structural and functional properties of the cells (Shapiro, 1995; Davey and Kell, 1996). As m a n y as 2000-3000 cells s ~can be analyzed with a standard flow cytometer, so that cytometric analysis is not only much faster than conventional epifluorescent analysis, but also more precise. A typical flow cytometric enumeration of bacteria will take 2-3 minutes, including sample handling and data analysis, compared to 15-30 minutes for a microscopic enumeration. In addition, the precision of routine cytometric enumerations is greater, with an average coefficient of variation of replicate samples of around 2% compared to the 10-25% that is often reported from microscopic counts (Kirchman, 1993; del Giorgio et al., 1996). In natural waters there are large numbers of particles and colloids within the range of cell sizes of bacterioplankton. It is impossible to distinguish natural bacteria from other submicron-sized particles on the basis of light scattering alone. For this reason, bacteria must be stained with a fluorochrome so that fluorescence can be used to discriminate cells from the multitude of particles of similar light scattering properties. The wide array of stains available for use in conjunction with flow cytometry (Table 8.2) provides ecologically valuable information about in situ bacterial cells.
Flow cytometers
Most current commercial flow cytometers share the same basic features and differ only in their capacity to sort and the light source used. Many commercial benchtop instruments are equipped with an argon laser with an excitation wavelength of 488 nm (e.g. the Becton Dickinson FACS line of flow cytometers/cell sorters, which are sufficiently portable and robust to be used on a ship or at a field site). Some cytometers, however, are equipped with mercury arc lamps (i.e. Skatron Argus flow cytometer), and a variety of other lasers are available, including He-Cd (emission from 320-440 nm), He-Ne (emission at 633 nm), and diode lasers emitting at >650nm. Some instruments can accommodate two lasers, which extends the capability of the instrument. The choice of instrument and light source depends on the type of analysis to be performed, and in turn the choice of fluorochromes will depend on the available excitation wavelength. There is a variety of fluorochromes that covers the entire range of available excitation wavelengths, with the greatest diversity in blueexcitable fluorochromes (Table 8.2). Since most modern commercial cytometers are equipped with a 488 nm argon laser, we provide below a protocol for bacterial enumeration protocol based on a blue-excitation fluorochrome. 138
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Sample fixation It is not necessary to fix or permeabilize cells for cytometric analysis. Fresh, unpreserved samples may be enumerated with cell-permeant stains, such as SYTO 13. A number of protocols to assess cell activity in fact require live samples. For samples that cannot be processed immediately and must be stored, fixation with 1-2% glutaraldehyde is recommended, del Giorgio et al. (1996) and Troussellier et al. (1995) found that fixation with formalin decreased cell fluorescence relative to glutaraldehyde or paraformaldehyde. Fixed samples should be stored frozen, optimally in liquid nitrogen.
Stains There is a wide variety of fluorescent stains that can be used in conjunction with flow cytometry to enumerate bacteria. Early cytometric counts of natural bacteria utilized UV-excitable nucleic acid stains such as DAPI (Button and Robertson, 1993) and Hoescht 33342 (Monger and Landry, 1993). But as mentioned above, most commercial cytometers are now equipped with an argon laser as the light source, and therefore there is increasing use of green-fluorescing nucleic acid stains that have excitation maxima at or close to the 488 nm laser. Most of these are manufactured by Molecular Probes (see Supplies section), including the YOYO, YOPRO, SYBR, SYTO and TOPRO families of nucleic acid stains (del Giorgio et al., 1996; Marie et al., 1996; LeBaron et al., 1998a). Each family of fluorochromes is composed of structurally similar compounds that differ slightly in the maximum excitation and emission wavelengths, and this allows the user to choose the one that best fits the application and the available instrumentation. For bacterial enumeration in a regular argon-based flow cytometer, SYTO 13 is one of the most widely used fluorochromes (del Giorgio et al., 1996; Gasol and del Giorgio, 2000). The SYTO family of stains is characterized by ahnost complete permeability to cell membranes, therefore no permeabilization step is required to attain effective staining of live or dead cells. The stain SYBR-II can also be used with live cells (Lebaron et al., 1998a). Other fluorochromes, such as TOPRO 1, also match the argon laser emission and have good spectral qualities, but require permeabilization of cells with detergent (Li et al. 1995). Pretreatment include buffers (acting also as cell permeants), such as Triton X-100 (Button and Robertson, 1993, Li et al., 1995), TE buffer (Marie et al., 1996), EDTA or EGTA (Kaprelyants and Kell, 1992; L6pez-Amor6s et al., 1995). Most blueexcitation nucleic acid stains require short incubation times, generally less than 15 minutes (del Giorgio et al., 1996; Marie et al., 1996; Veldhuis et al., 1997). Some authors have successfully counted bacteria without any pretreatments such as those discussed here; it is up to each researcher to decide whether pre-treatment is necessary. Since detergents may elevate the sample background, it is convenient to minimize their use.
Counting Enumerating bacteria by flow cytomety implies detecting stained bacterial cells in a given volume of sample processed by the cytometer. Only a 140
few commercial cytometers, however (e.g. the Coulter XL and the Ortho Cytoron Absolute) are equipped to precisely measure the volume of sample processed. Most flow cytometers have fixed flow rate settings that only roughly approximate the true flow rate. In addition, the flow rate at a fixed setting will vary due to electronic and hardware variations. There are at least three ways of obtaining absolute counts under these conditions: (1) a known amount of reference beads can be added to each sample (Cantineaux et al., 1993); (2) the flow can be calibrated during each work day; or (3) each sample can be weighed before and after the run. The last alternative is time consuming, and in addition, it may be less accurate, because there may be backflow of sheath fluid into the tube, which confounds the actual amount of sample volume processed. Alternative 2 (daily calibration) gives good results but requires a very stable instrument (Gasol and del Giorgio, 2000). Calibration of the flow can be done easily by weighing a tube containing water, processing various volumes through the cytometer, estimating the time needed for each volume to go through, and reweighing the tube. Most researchers, however, use alternative 1 (reference beads). This approach is accurate, fast and, in addition to allowing quantitative determination of cell abundance, also provides an internal standard that can be used to assess instrument preformance and to standardize scatter and fluorescence measurements for quantitative applications. In the latter application, the average channel value for the bead population is used to normalize signals for each parameter (scatter, fluorescence) of each bacterial population.
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A variety of green-fluorescent microbeads can be used as internal standards (i.e. Fluoresbrite, Polysciences, or FluoSpheres, Molecular Probes, see Supplies section). Beads ranging in size from 0.75 to 1.0 txm in diameter usually provide the least overlap with the natural bacterial populations and are thus most effective for enumeration and standardization. A stock solution of 10'~beads is prepared with 0.1-tim-filtered distilled water. Final concentrations of 3c/~ ethanol, or 0.5% glutaraldehyde or formaldehyde, can be added to the stock solution to prevent bacterial growth. The bead stock is dispensed into each sample to yield a final bead density about 1-10% of the expected density of target cells. For a bacterioplankton sample with an abundance of 10" cells ml ', a final bead density of 1-5 x 10 beads ml 'is appropriate. The bead density in the stock solution must be carefully assessed on a routine basis. Larger beads (> 2.0 t~m) can be counted in a Coulter particle analyzer, but this method is less effective for the smaller beads which are generally used for bacterial work. Alternatively, bead density can be determined using regular epifluorescence microscopy, but this is time consuming and not particularly precise. A more effective approach is to use a primary reference bead solution where the bead density is precisely known (e.g. TrueCount, Becton Dickinson), and to compare this to the working bead solution using the flow cytometer. 141
Cytometric protocol
Materials required Equipment Flow cytometer Pipettors for quantitative delivery of water samples, reference beads, and fluorochromes • Vortex mixer • -20°C freezer for storage of samples •
•
Supplies • • • • •
Cytometer tubes. Some instruments, e.g. the Becton Dickinson FACSCalibur, require the use of a specific type of tube Sheath fluid Reference beads: 0.75-1.0pro green fluorescing beads, FluoSpheres (Molecular Probes) or similar DMSO to dilute fluorochromes Primary bead reference solution to determine bead working stock density (TrueCount, Becton Dickinson).
Solutions •
SYTO 13 stock solution, 0.5 mM in DMSO.The SYTO dyes, as well as most nucleic acid stains sold by Molecular Probes, are delivered as a I-5 mM solution in DMSO. This solution is diluted with high-grade DMSO to a final concentration of 0.5 mM. This working solution can be distributed among several microfuge vials, which are then stored at -20°C, so that only a portion need be thawed at any given time. The working SYTO 13 solution can be stored for months this way. • Sheath fluid. Commercial sheath fluid can be purchased as a weak NaCI solution containing various buffers and antimicrobial agents (i.e. FACSFIow, Becton Dickinson). For most bacterial applications, however, 0.2-pro-filtered distilled water can be used as sheath fluid. For some applications, including marine samples, or when the sample is sorted into the sheath fluid, a weak (I-5%) NaCI solution, also filtered through 0.2 pro, may be more appropriate than distilled water. 0.2-~m-filtered seawater has also been recommended for marine samples (Olson et al., 1993). Bacterial growth may occur in filteredsterilized water, so it is recommended that either the water be filtered immediately prior to filling the sheath fluid tank, or autoclaved before storage. • Formaldehyde or glutaraldehyde for sample fixation,0.2 pm flltered.'See page 140.'
Cytometric analysis of samples The following is a basic cytometric protocol using a standard b e n c h t o p i n s t r u m e n t (FACSCalibur, Becton Dickinson) e q u i p p e d with a 488 n m argon laser, and the cell-permeant nucleic acid stain SYTO 13. There are multiple variations to this protocol, d e p e n d i n g on the available instrum e n t and the actual application, but the basic steps outlined below are c o m m o n to m o s t protocols.
142
1. Fix sample if necessary and store fixed sample at -20°C or below if it cannot be analyzed immediately. If samples have large amounts of particulate detritus and phytoplankton, it is advisable to pre-filter the water through 20 to 50 txm mesh netting to avoid clogging the fluidics system. 2. Pipette 0.5 ml of fresh or fixed sample into a cytometer tube. 3. Pipette sufficient v o l u m e of reference bead solution to attain a final bead concentration in the sample in the range 0.5- 5 x 10 ~beads ml '. 4. Pipette 2 ~1 of 0.5 mM SYTO 13 working solution into the tube, for a final sample concentration of 0.2 pM SYTO. The final concentration of SYTO 13 is not critical, and a range of concentrations from < 1 I_IM to > 5 lJM have been used (del Giorgio et al., 1996; Lebaron et al., 1998a). If another type of nucleic acid stain is used, for example TOPRO, bacteria m a y have to be permeabilized prior to staining. 5. Vortex each tube for 5 s, and leave for 5-10 rain in the dark. The sample is n o w ready to be run in the flow cytometer. 6. The detection of cell populations in the cytometer requires empirically adjusting the voltage and gain of the Photomultiplier Tubes (PMT) for side scatter and fluorescence, and the gain of the forward scatter photodiode, so that the population of interest is clearly visible. As a general rule, samples for bacterial enumeration are analyzed in logarithmic mode, and bacteria are detected using a combination of side scatter and green fluorescence (emitted by the blue-excitation nucleic acid fluorochrome used to stain cells). In lnost instruments, forward scatter provides less resolution than side scatter. In logarithmic acquisition m o d e only the PMT voltage for side scatter can be adjusted, in addition to the gain for the forward scatter diode. To establish the PMT voltages for the flow cytometer, it is convenient to begin with one or several water samples with reference beads added but no stain. These samples are used to adjust the photomultiplier voltage for side scatter and green fluorescence to allow detection of beads and to minimize background. This background often appears as a rectangular area with a large n u m b e r of events near the origin in the fluorescence and scatter axes. It results from a combination of electronic noise and weakly fluorescent particles or colloids in the sample, which may include cell remnants and viruses. The background can be reduced by increasing the threshold of green fluorescence necessary to trigger an event to a level where most of these non-bacterial events do not appear. SYTO 13 is then a d d e d to these samples, and the stained bacteria will now appear as a fairly dispersed 'cloud' in the cytogram, often consisting of several distinct populations (e.g. high and low DNA-content cells, Figure 8.1). The voltages are further adjusted so that the entire bacterial 'cloud' is in the center of the cytogram defined by side scatter and green fluorescence, the reference beads also appear in the plane, and the background noise, if any, appears compressed against the axes (see Figure 8.1). 7. Once an appropriate set of PMT voltages have been established, they should be saved and reused for all subsquent analyses of similar samples in the same instrument. There will be minor instrument 143
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variations f r o m d a y to d a y w h i c h can be corrected b y use of the internal b e a d reference. 8. The stained bacterial s a m p l e s can n o w be r u n in the cytometer. It is c o n v e n i e n t to use the lowest possible flow rate, a n d not to exceed a rate of 2000 events s ' (preferably 1000 events s r) p a s s i n g the laser beam. H i g h e v e n t rates greatly increase the possibility of h a v i n g d o u b l e events and cell coincidence, w i t h potentially significant bias in the cell count. If at the lowest possible flow rate setting, the e v e n t rate is still over 2000 s ', w h i c h is c o m m o n in s a m p l e s w i t h m o r e than 6 x 10" cells ml ', it is preferable to dilute the s a m p l e w i t h 0.1-pro-filtered distilled w a t e r or w e a k saline solution just prior to the analysis. If s a m p l e s are diluted after the stain a n d the b e a d s t a n d a r d has b e e n a d d e d , then no correction is required in s u b s e q u e n t calculations b e c a u s e the original b e a d to cell ratio is retained. 9. A c q u i r e at least 15 000 events and save the resulting list-mode data file for later analysis. This will take a b o u t 5 to 40 s.
144
10. Once all the samples have been run in the cytometer, the data files can be analyzed using a variety of software packages, usually included with the instrument. The data are typically analyzed in a plot of green fluorescence versus side scatter, as in Figure 8.1. Windows are drawn around the bead and bacterial populations, and the number of events in each window are recorded. Bacterial cell abundance is derived as follows: Bacterial density (cells ml ') = [(number of cells / number of beads) x bead density (beads ml ~)] x Dilution Factor (fixative + bead solution added)
Cytometric determination of cell size Light scattering at different angles is related to a wide range of cellular characteristics, but scattering at small angles is mostly a function of particle volume and secondarily of particle shape (Button and Roberston, 1993; Allman et al., 1990). Koch et al. (1996) outlined the theoretical basis of the relationship between cell size and forward scatter; this relationship has been used to assess bacterioplankton cell size (Robertson et al., 1998). A theoretical algorithm derived from light scattering theory for particles of the size range of bacteria, which must be calibrated for each type of instrument, is used to predict size as a nonlinear function of cell scatter. The algorithm works well with empirical relationships between forward scatter and size of bacterial cultures and beads, but has yet to be shown effective in the size range of natural bacterioplankton. An alternative to using scattered light as an index of bacterial size is the use of the cellspecific fluorescence of DNA-bound stains. Veldhuis et al. (1997) found that DNA content, as estimated with PicoGreen, varies with cellular C and N content, at least for picoalgae and nanoalgae. There is a relatively good relationship between image analysis measurements of planktonic bacterial size (in the range 0.03-0.09 lam ~) and the average green SYTO 13 fluorescence per cell (Gasol and del Giorgio, 2000). This observation suggests that DNA-related fluorescence can be used as a surrogate for bacterial size, although calibration using image analysis is needed because there is little theoretical basis for the relationship. Regardless of whether scatter or fluorescence is used to estimate cell size, it is necessary to determine the mean cell size using image analysis in a subset of the samples and empirically establish the relationship with the cytometric parameters.
Co-enumeration of heterotrophic and autotrophic bacteria It is possible to discriminate and enumerate autotrophic and heterotrophic prokaryotes in the same water sample (Olson et al., 1993). An important group of oceanic phytoplankton, bacteria containing chlorophyll-a, or prochlorophytes, are most effectively enumerated via flow cytometry (Chisholm et al., 1988; Campbell et al., 1994; Sieracki et al., 1995; Campbell, Chapter 16, this volume). The argon laser can be used to selectively detect autotrophic and heterotrophic bacteria because photopigment emissions are sufficiently distinct from the emission of most blue-excitable stains. The protocol is essentially the same as for hetero145
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trophic bacteria, but red a n d / o r orange fluorescence is used in addition to green fluorescence and side scatter to discriminate autotrophic prokaryotes. The principle is that all cells will fluoresce green after staining with SYTO-13 (or similar nucleic acid stain), but autotrophic cells will in addition show significant red (or orange) fluorescence due to photosynthetic pigments. The total cell density is obtained from a cytogram of green fluorescence versus side scatter, as described above. Details of the cytometric analysis of autotrophic bacteria and phytoplankton appear in Chapter 30 of this volume.
Cell-specific indices of cell activity At present, there is considerable controversy concerning what fractions of in situ bacterial cells, enumerated by standard epifluorescence staining methods, are metabolically active and growing, dormant (in a state of starvation survival, Morita, 1997), or actually dead. Williams et al. (1998) suggested that marine bacteria can be categorized as dead (cells with damaged membranes), live but inactive (intact membrane but not detectable with a universal 16S rRNA oligonucleotide probe), or live and active (intact membrane, detectable with a rRNA probe). We concluded from analysis of the CTC assay (B. Sherr et al., 1999) that CTC+ cells probably have the highest level of metabolic activity in a bacteria assemblage. Based on cell-specific DNA content and membrane integrity, Gasol et al. (1999) proposed that in situ bacteria can be sorted into five groups: (1) cell fragments or ghosts; (2) dead cells; (3) live but inactive cells; (4) slowgrowing cells; and (5) large bacteria growing at fast rates. Here we present a protocol for one approach to determination of cell-specific metabolic activity in living cells: the CTC assay. Other approaches to determine single-cell physiological state are outlined in Chapter 10 of this volume.
ETS-active cells using CTC The fluorogenic redox dye 5-cyano-2,3-ditolyl tetrazolium chloride (CTC) has been used in both freshwater and marine systems as a vital stain for enumeration of actively respiring bacteria in situ (del Giorgio and Scarborough, 1995; del Giorgio et al., 1997; B. Sherr et al., 1999). CTC is water soluble and non-fluorescent in its oxidized state, but becomes highly fluorescent and insoluble when reduced (Rodriguez et al., 1992). The CTC method has been criticized as yielding underestimates of the proportion of active cells in bacterial assemblages. We have addressed these criticisms and have made a case for interpreting CTC-positive (CTC+) cells as the most highly active cells in the assemblage (B. Sherr et al., 1999). Changes in the proportion of CTC+ cells can signal changes in the overall metabolic activity of in situ bacteria (Choi et aI., 1999; B. Sherr et al., 1999; E. Sherr et al., 1999). The abundance of CTC+ cells can be enumerated via epifluorescence microscopy or flow cytometry; flow cytometry is more sensitive and yields higher counts of CTC+ cells compared to microscopy (B. Sherr et al., 1999; Sieracki et al., 1999).
146
Materials required Equipment •
Epifluorescence microscope outfitted with UV and blue light filter sets. Although the fluorescence of reduced CTC is most intense under green light illumination, we have found that CTC+ cells are easily observed under blue light illumination. • Constant temperature dark incubator at in situ temperature • Adjustable pipettors: 10-1000 HI, 10-200 lul • Low temperature freezer (-20°C or lower) for storing preserved CTC samples
Supplies • • •
2 ml cryovials, or plastic vials that can be stored at low temperature Ziploc or other brand of plastic freezer bags CTC powder - - from Polysciences Inc., catalogue # 19292, I gram = $409
Solutions •
50 mM CTC solution. Note: the CTC reagent is a potent metabolic poison; when making up the solution and filtering it through the 0.2 lam acrodisc, use gloves and carefully discard the used acrodisc.Wash hands well after handling CTC. Dissolve 100 mg CTC powder in 6.6 ml of deionized water, filter through a 0.2/am Acrodisc filtration unit (Millipore) to remove any remaining undissolved CTC, store at 4-5°C in a dark glass vial, e.g. a 20 ml glass scintillation vial covered in black electrical tape. For 0.9 ml seawater samples, this amount of CTC solution would be enough for about 60 assays.
Recent batches of CTC we have obtained from Polysciences have dissolved readily in deionized water without need for extensive stirring or sonication. In our experience, the prepared CTC solution is good for weeks to a couple of m o n t h s if stored at 4-5°C and kept out of direct light. Notes: CTC reagent should be stored in the refrigerator in a dark container. DO NOT FREEZE either the CTC p o w d e r or the stock solution; freezing denatures the compound! Each batch of CTC p o w d e r should be tested immediately. Make up at least a small a m o u n t of CTC solution and test with bacteria k n o w n to be actively growing. If there is not a strong positive reaction to the CTC solution (abundant bright red cells) then the CTC p o w d e r m a y be bad; the c o m p a n y can be persuaded to replace it. •
Sodium-borate buffered formalin, prepare as described in the epifluorescence microscopy section
Sampling Take a water sample using a clean, rinsed, 'live' bottle, i.e. one that has not had any fixative in it. To economize o51 CTC, we usually use 0.9 ml subsamples; but for samples with low bacterial abundance, e.g. less than 3 5 x 10"cells m l , the sample a m o u n t can be increased to 1.8 ml and a larger incubation vial used. 147
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Procedure Note: try to minimize light of wavelengths < 420 nm during handling of CTC and samples; direct sunlight should be completely avoided. 1. For each sample, dispense two 0.9 ml (or 1.8 ml) subsamples into two labeled cryovials. Since CTC is a vital stain that is reduced only by living cells, avoid contamination of the samples with fixatives or other toxic substances. 2. Add 100 gl of CTC solution to each 0.9 ml sample (1:10 dilution, 5 mM CTC final concentration) (or 200 ~tl to each 1.8 ml sample) and gently mix. In this assay, the concentration of CTC is much higher than used for passively binding stains such as DAPI because dissolved CTC must be actively reduced to the fluorogenic compound by the bacterial ETS. 3. Incubate IN THE DARK for 2-3h at in situ temperature. We have found 3 h to be sufficient for most systems we have examined, but incubation times of up to 8h have been used (del Giorgio and Scarborough, 1995). Note: when analysis is via flow cytometry rather than microscopic inspection, optimum CTC concentration may be lower, e.g. 2.5 mM rather than 5 mM, and optimum incubation times may be shorter, e.g. < 1 h (B. Sherr et al., 1999; Sieracki et al., 1999). For individual systems, one should run control experiments over a range of CTC concentrations (0.5 to 5 mM) and incubation times (0.5-6 h) to determine optimum conditions for determining CTC+ cells. The abundance of CTC+ cells will reach a plateau when the concentration of CTC reagent and the time of incubation are optimized. 4. Add 5% final concentration 0.2-p_m-filtered formalin (50 ~1 for 1 ml sample or 100gl for 2ml samples), shake or vortex to mix. Immediately freeze vials in liquid nitrogen (or in a -20°C freezer if liquid nitrogen is unavailable) in labeled and dated Zip-lock bags (one bag for each set of CTC samples). Keep frozen until analyzed. If transport is necessary, frozen CTC samples should be shipped with dry ice.
Analysis of CTC+ samples usingepifluorescence microscopy 1. Thaw sample, add 25 ~tl ml ' of sample of 200 gg mU DAPI solution to
cryovial, shake or vortex to mix, incubate for about 10 min in the dark. 2. Filter entire sample onto a 0.2 ~tm black filter, mount on a slide and
immediately inspect via epifluorescence microscopy. 3. Identify bacterial cells with the UV light filter set (cells stained with
DAPI), and then switch to the blue light filter set to see whether the cells have accumulated reduced CTC (they will look reddish-orange). Be careful to avoid counting 'false positives', i.e. particulate CTC not associated with DAPI-stained bacterial cells. The photopigments of coccoid cyanobacteria have a fluorescence similar to that of CTC. In samples with cyanobacteria, one should prepare control samples without CTC in order to estimate the abundance of cyanobactera. With practice, it is usually possible to visually distinguish between cyanobacteria and CTC+ bacteria. 148
Analysis of CTC+ samplesusing flow cytometry CTC+ ceils can be easily enumerated using flow cytometry (del Giorgio et al., 1997; Sieracki et al., 1999). Sample preparation is the same as above. 1. Thaw samples or use fresh samples immediately after incubation. Place 0.5 ml of sample in a cytometer tube. Add 1.0 lJm diameter green reference beads (as used for total bacterial enumeration) to yield a final concentration of 0.5-2 x 105 ml 1. 2. Determine the appropriate PMT voltages. CTC+ cells are detected using a combination of either orange or red fluorescence and side light scatter. The background can be eliminated by setting the threshold in red fluorescence. The presence of autotrophic picoplankton in samples may interfere with the enumeration of CTC+ cells, because fluorescence of chlorophyll and phycobiliprotein pigments is similar to that of CTC. Although autotrophic picoplankton are generally larger than heterotrophic bacteria and can usually be discriminated on the basis of light scatter, this may not always be true. If samples are suspected of having a high density of autotrophic picoplankton, it may be necessary to enumerate these cells in an unstained aliquot of the sample, and then substract the number from the CTC+ counts. 3. Run the sample at the lowest flow rate possible, and acquire at least 10 000 events. 4. After CTC-positive cells have been enumerated, the total number of bacteria in the sample can be determined by adding SYTO 13 to the same cytometer tube and following the protocol for total bacteria described above. SYTO 13 and CTC-stained cells fluorescence differently and in theory one could enumerate the total and the CTC+ cells in the same double-stained sample. In practice, however, it is often difficult to fully separate CTC+ from CTC- cells in a sample double stained with SYTO 13. It is for this reason that we suggest that the sample first be run with CTC alone, then double stained with SYTO and run again to obtain the total count, which includes both CTC+ and CTC- cells. 5. Data analysis proceeds as for total enumeration as described above, except that data are analysed in a plot of red (or orange) fluorescence versus side scatter, as shown in Figure 8.2. The mean cell fluorescence may be linked to mean cell activity and may be useful to complement the counts of CTC+ cells (B. Sherr et al., 1999).
RESULTS The abundance of heterotrophic bacteria has been analyzed in a wide variety of aquatic ecosystems. Across systems of differing trophic states, bacterial abundances are positively related to phytoplankton biomass, and are generally in the range 0.5 × 10' to 5 x 10 '¢' cells 1 ' (Bird and Kalff, 1984; Cole et al., 1988). The proportion of bacterial biomass to phytoplankton biomass increases with decreasing phytoplankton standing 149
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stock and system productivity; in oligotrophic systems, bacteria can be a large (>50% of total) c o m p o n e n t of plankton biomass (Bird and Kalff, 1984; Cole et al., 1988; Fuhrman et al., 1989; Cho and Azam, 1988, 1990; Simon et al., 1992; Gasol et al., 1997). The range in bacterial activity tends to be much greater than the range in bacterial abundance (Cole et al., 1988; Ducklow and Carlson, 1992). For example, in recent work off the Oregon coast, bacterial cell-specific [~H]leucine incorporation rates varied > 70-fold, while bacterial abundance varied only -10-fold (E. Sherr et al., 1999). Such observations show that there can be a wide variation in the average cell-specific activity levels among bacterial assemblages. The CTC assay is one approach to determining, at least qualitatively, the distribution of cell-specific metabolic activity in natural assemblages of bacteria. The proportion of bacterial cells in situ that reduce detectable quantities of CTC, as a measure of active ETS, is generally < 10% (Gasol et al., 1995; del Giorgio and Scarborough, 1995; Karner and Fuhrman, 1997;
150
B. Sherr et al., 1999). We are aware of, and have addressed, criticisms of the CTC assay as an index of active cells (B. Sherr et al., 1999). Some of these criticisms are unfounded. For example, the idea that not all aerobic, organotrophic bacteria in the sea can reduce CTC has little basis. Strong CTC reduction during active growth was found for each of 27 bacterial cellular clones, representing a wide range of phylogenetic groups, isolated from Oregon shelf seawater (B. Sherr et al., 1999). In addition, tip to 80-90% of total bacterioplankton cells in seawater could be induced to become CTC+ (Choi et al., 1999). We' are not aware of any aerobic, heterotrophic marine bacterium shown to be incapable of detectable CTC reduction during active growth. Toxicity of CTC or of its formazan precipitate to bacterial cells has also been a criticism of the technique (Ullrich eta/., 1996, Karner and Fuhrman, 1997). CTC toxicity, if a problem, does not appear to have an immediate effect, del Giorgio et al. (1997) used flow cytometry to measure the intracellular accumulation of CT-formazan and noted that mean red fluorescence emission per cell continued to increase for several hours after the number of CTC+ cells had stabilized, indicating that cells continued to reduce CTC and accumulate formazan long after they were visibly stained under flow cytometry. This result suggests that cells are not killed instantly, but rather continue to function for hours after exposure to CTC. Results of both laboratory and field studies support the idea that reduction of sufficient CTC for a cell to be scored as positive is a characteristic of actively growing cells (Pyle et al., 1995; Choi et al., 1996, 1999; Smith, 1998; B. Sherr et al., 1999; E. Sherr et al., 1999; Sieracki et al., 1999).
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DISCUSSION AND FUTURE DIRECTIONS Bacterial abundance, cell size, biomass, and activity are among the most fundamental properties that characterize natural aquatic systems, and the measurement of these parameters is central to aquatic microbial ecology. Epifluorescence techiques are now routinely performed in most microbial laboratories, and image analysis has increasingly been used to determine cell size and other cellular properties. The application of flow cytometry to the determination of the abundance and biomass of natural bacterial assemblages is still not routine, but will without doubt continue to increase as instruments and techniques become more available. Flow cytometry offers the possibility of assessing not only total abundance with great speed and precision, but also properties of individual cells, such as size and metabolic activity. Cytometry also offers the possibility of physically separating, via cell sorting, subpopulations based on various cellular properties. This is an application that will be increasingly used in microbial ecology. Flow cytometry and image analysis (Viles and Sieracki, 1992; Sieracki and Viles, 1998), combined with selective staining methods, permit determination of physiological condition of ilz situ bacterioplankton. Physiological characteristics that can now be detected include: cell-specific 151
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content of DNA (Li et aI., 1995; Veldhuis et al., 1997, Gasol et al. 1999), rRNA (Kramer and Singleton, 1993; Lee and Kemp, 1994; Binder and Liu, 1998), or protein (Zubkov et al., 1999); polarization state of the cell membrane (Mason et al. 1995, Nebe-Von Caron et al. 1998); damaged versus intact cell membranes (Williams et al., 1998; Lebaron et al., 1998b), and level of activity of electron transport systems (Gasol et al. 1995; del Giorgio et al., 1997; B. Sherr et al., 1999). In addition, fluorescent in situ hybridization (FISH) using targeted 16s rRNA probes allows enumeration of specific phylogenetic groups of aquatic bacteria (Amann et al., 1990a and b, Ouverney and Fuhrman, 1999; Simek et al., 1999). Two recent studies combining flow cytometry and specific staining methods serve as examples of the future direction of bacterial enumeration. Servais et al. (1999) found, using flow cytometric sorting of radiolabeled bacteria from a eutrophic mesocosm experiment, that larger cells (> 0.25 ~tm3) had on average 10-fold higher leucine incorporation rates than smaller bacterial cells (< 0.25 ~tm~). Gasol et al. (1999) reported that the numbers of bacteria in Mediterranean seawater that could be classified as living (with an intact cell membrane), having an organized nucleoid region, and having high DNA-content were equivalent and about 60-70% of total cells. These studies demonstrate that advances in bacterial enumeration open the possibility of addressing questions that until recently were beyond the realm of technical feasiblility. In the future, the combination of molecular techniques with image analysis and flow cytometry will allow routine and efficient determination of the physiological and phylogenetic structure of bacterial assemblages at the same time as total abundance, cell size and biomass are measured.
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9 Isolation of Oligobacteria D o n K Button', Betsy R Robertson 2 and P h a m Q u a n g 3 'Institute of Marine Scienceand Department of Chemistry and Biochemistry,University of Alaska, Fairbanks,Alaska 9 9 775, USA;~lnstitute of Marine Science,University of Alaska, Fairbanks,Alaska, USA; 3Department of Mathematical Sciences,University of Alaska, Fairbanks,Alaska, USA
CONTENTS Introduction Nomenclature Method I. Spread plate isolation Method 2. Extinction culture isolation Properties of extinction cultures and future directions Conclusion
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Isolation of typical marine bacteria is of value because the resulting cultures provide material essential for understanding bacterioplankton. These prokaryotes constitute a main component of aquatic biomass and regulate the concentrations of dissolved organic carbon, yet few examples have been cultivated and significant uncertainty surrounds their properties. Marine bacteria were first recognized in the 18th century (Zobell and Upham, 1944) and many species able to reproduce at the large nutrient concentrations used in enrichment culture have since been described. The conventional techniques used produce cultures from only about 0.1 ~7, of the organisms present. However, the high return rates of carbon dioxide from radiolabeled glucose (Andrews and Williams, 1971) and hydrocarbons (Robertson et al., 1973) suggested activity that is consistent only with larger populations, an abundance first observed by staining with fluorescent dyes (Daley and Hobbie, 1975). These populations have been characterized by electron microscopy (Schmidt et al., 1991), epifluorescence image analysis (Sieracki and Viles, 1992), flow cytometry (Robertson et al., 1998), and various molecular probes. It appears that most marine bacterial phylotypes and ecotypes are yet to be cultivated. Evidence includes the low success in isolation, the presence of unrecognized DNA sequences in seawater (Britshgi and Giovannoni, 1991) from bacterium-sized organisms, the absence of the 1-3 kb genome organisms among marine isolates (Cammack, 1997), and the 50-fold increase in METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0-12 521530 4
C o p y r i g h t © 2001 A c a d e m i c Press Ltd All rights of reproduction ill anv form reserved
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specific affinity for radiolabeled substrate that is obtainable by brief warming of cold seawater. Reasons for the low success include difficulty in resuscitation a n d / o r perpetuation from the normal state, damage due to viral attack, and substrate-accelerated death (Whitesides and Oliver, 1997) due to unintentionally included substrates. There is also uncertainty about the physiological status of the small forms that are counted as bacteria, and commensal or symbiotic relationships among the pelagic forms. Some think m a n y particles counted as bacteria are incomplete cells (Gasol et al., 1999) and others find the fraction of small cells to be as active as the large (Button and Robertson, 2000). The frequency of isolation success has been increased by use of extinction cultures where viabilities can approach 10% even in oligotrophic low-productivity waters. The extinction culture technique is based on the observation that when bacteria are filtered from seawater and then re-inoculated into filtered seawater, the population will regenerate. The resulting populations were first called seawater cultures (Jannasch, 1979). When the inoculum is large, i.e. 10'~ unfiltered seawater, the rate of increase can be used to determine the rate of growth (Button and Robertson, 1993). When it is small, just a few organisms, it can be used in combination with the number of organisms inoculated to determine the culturability or 'viability' of those originally present (Robertson et al., 1998).
NOMENCLATURE Aquatic heterotrophic bacteria are included as the main component of the picoplankton, free living 0.2-2.0 Ixm diameter heterotrophic organisms comprised of species from the Eubacterial and the Archaeal kingdoms. Their ecophysiology is transthreptic (surface feeding), i.e. they transport organics across a membrane impermeable to polar substrates to effect internal concentrations sufficient for growth, and as such they comprise a major division of organisms that also includes smaller numbers of yeasts, molds, and actinomyces. Viability is taken as the fraction of observable organisms that grow to detectable populations. Oligobacteria are defined as bacteria that can accumulate dissolved organics from low concentrations. These concentrations are imprecisely defined and not well understood, particularly the important amphipathic components. But in terms of kinetics the ratio of the specific affinity constant to the affinity constant (a ° ~/K,) should be large, of the order of log 1000 to give an oligotrophic capacity >3 (Button, 1998) to supply small amounts of cytoplasm with substrate by way of large numbers of permeases. Since chemical contamination is presently unavoidable and can exceed the amounts added (and the analyses are difficult), background substrates remain of concern. Defined monomers are in the range Fig 1 ' or less. Confounding the issue are substrate inhibition and stimulation by small concentrations of substrate, in combination with rising temperatures that may trigger emergence from a resting state. It is likely that a range of facultative to obligate oligotrophs exists and that the true oligotrophs are harder to cultivate. It 162
is also likely that true oligotrophs have higher specific affinities for substrate than those isolated to date because of the more modest requirements for regulator genes and their energy and space-consuming protein products (Button and Robertson, 2000). Organisrns in culture collections are generally comprised of facultative oligotrophs and copeotrophs such as Escherichia coil and Staphylococcus aureus. Oligobacteria are smaller than copeotrophs giving rise to the term ultramicrobacteria (Torrella and Morita, 1981). They are low in solids content and DNA (Button and Robertson, 2000), probably as a way of maximizing use of their energy reserves for growth and increasing their surface to volume ratio to maximize their specific affinities for nutrients.
METHOD
I. S P R E A D P L A T E I S O L A T I O N
An early potential, but incompletely developed, technique used to isolate marine bacteria is to select by microscope microcolonies formed on agar spread plates with minimal or no organic nutrient additions. In a preliminary experiment, we incubated agar plates prepared without added nutrients for about 5 months. The glass plates were nearly filled with 1.5% agar and incubated in a cut-off 20 1 carboy with a little water in the bottom along with a wick of paper towels to keep the air moist. The whole apparatus was closed with a glass plate sealed w,ith stop-cock grease and incubated at 10°C. Numerous small colonies appeared, more than expected from experiences with traditional seawater plating techniques, but additional observations were not made. Agar has components useful to many bacteria and modifications of the technique involve the use of glass fiber filters floated over seawater contained in depressions formed in glass culture plates, with incubations in lmmidified containers. Colonies were observed by microscopy and selected from the filter surface. These are promising but incompletely developed techniques.
M E T H O D 2. E X T I N C T I O N ISOLATION
CULTURE
To minimize inadvertent addition of organics and the effects of solid surfaces associated with culture plates, organisms can be isolated by dilution to extinction. A total population count is required to calculate the appropriate dilution. This can be easily done, for example aboard ship, by epifluorescence microscopy. Confirmation by flow cytometrv (Robertson et al., 1098) on preserved samples is useful where accurate population values are of interest because in oligotrophic waters there is significant subjectivity used in deciding if dim particles are srnallgenome bacteria, condensed nucleoids, bacteria that have lost some DNA, or large viruses. 163
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Population determination Samples are treated with a mixture of formaldehyde (to 0.5%), Triton X-100 (0.1%) and DAPI (0.5 lJg m1-1) for three hours at room temperature. This results in brighter staining than 1 h 10~-'C conditions used for DNA quantitation by flow cytometry. For microscopy the formaldehyde m a y be replaced with glutaraldehyde at 0.5%. A black polycarbonate 25 m m diameter 0.1 Bm pore-size filter is placed shiny side up on the glass frit of a vacuum filtration unit, and 1-3 ml of seawater is filtered through it. The wash is 1 ml of filtered seawater and is used also as a blank. A drop of immersion oil is spread onto a microscope slide, the filter applied, topped with another drop of oil and flattened under a cover slip, and the collected organisms viewed by epifluorescence microscopy with ocular grid. Excitation is in the UV with observation through a 430nm long-pass filter. This typically gives a population of near 50 organisms per 0.01mm 2 grid, and 300 are counted. For a 17 m m diameter frit the population ml ~ is the number of bacteria per g r i d / v o l u m e filtered x 22 000.
Viability calculations Values for viability or culturability are useful to determine the fraction of the population present that is capable of growth under conditions presented. Viability m a y be estimated (Quang et al., 1998) from
V-
In(l-p) X
(9.1)
The asymptotic standard error in viability A S E ( V ) is given by
ASE(V)-
1
'
p
X \, n(1- p)
(9.2)
The asymptotic standard error m a y be used to set an approximate 95~ confidence interval for true viability in the form V + 1.96 ASE(V). The coefficient of variation in the viability is C V ( V ) = A S E ( V ) / V where V
z n p X u
viability = number of vessels showing growth = number of vessels z/n = number of counted organisms inoculated into each vessel = estimated number of vessels of pure cultures.
=
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Pure culture production The n u m b e r of pure cultures expected to arise in the absence of allelopathic or synergistic interactions is given by u = -n(1 - p) ln(1 - p)
(9.3)
ASE(u) = ln(1 - p) + lx np(1 - p )
(9.4)
and the standard error by
The optimal inoculating population for minimal error in the determination of viability, X = 1.6/V, is given in Table 9.1. For example, if the culture viability is 10% then the vessels should be inoculated with 16 organisms from the original seawater sample using the population count as a base to calculate the dilution.
Table 9. I Inoculating population for minimal error in viability Viability (%) Population for minimal error 160 80 40 16 8 2
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The n u m b e r of pure cultures to be expected, with respect to viability, is s h o w n in Table 9.2. For example, at a viability of 8 ~ an average of 3.6 from 10 vessels should show growth and 3.5 of them are likely to contain pure cultures. By inoculating 30 vessels three times as m a n y pure cultures are expected and the error in determining viability is only about half as large. By increasing the inoculum to 30 organisms per vessel, only half as m a n y pure cultures are expected and the certainty of that n u m b e r is only within 2.2 of the 6.5 pure cultures expected within a confidence level of 95%.
Viability when the n u m b e r of species is known To improve the viability estimate, data for the n u m b e r of species observed can be added. For example, in a recent seawater sample from Resurrection Bay, Alaska, the species count from a series of 10 vessels inoculated with 10 organisms each was 2 3 0 5 4 5 1 0 6 1. Note that the n u m b e r of species may not be identified across the vessels and some m a y be redundant. Results from a second set was 6 4 5 5 5 4 4 6 6 2. in the uninoculated control the results were 0 0 4 0 4 4 3 2. The likelihood L,,,, (Rice, 1995) of observing
165
o
Table 9.2 The expected number of total and pure cultures at various values of viability for particular numbers of trials Viability (%) 1 2 4 8 12 20
Standard error (V) (%)
Vessels to show growth (number)
Pure cultures (number)
10 organisms inoculated into each of 10 vessels 0.9 1.0 0.9 1.2 1.8 1.8 1.5 3.3 2.7 1.6 3.6 3.5 1.5 7.0 3.6 1.1 8.6 2.7
Standard error for purity 0.8 1.0 0.9 0.3 0.3 1.1
2 4 8 16 20
10 organisms inoculated into each of 30 vessels 0.5 2.9 2.7 0.7 5.4 4.9 0.9 9.9 8.0 0.9 16.5 10.8 0.7 23.9 9.7 0.6 25.9 8.1
1.7 1.5 0.5 0.5 1.8
1 2 4 8 12 20
30 organisms inoculated into each of 10 vessels 0.5 2.6 2.2 0.5 4.5 3.3 0.5 7.0 3.6 0.3 9.1 2.2 0.2 9.7 1.0 0.1 10.0 0.1
1.0 0.6 0.3 1.2 1.3 0.7
1
1
2 4 8 12 20
30 organisms inoculated into each of 30 vessels 0.3 7.8 6.7 0.4 13.5 9.9 0.3 21.0 10.8 0.2 27.3 6.5 0.1 29.2 3.0 0.0 29.9 0.4
1.4
1.7
1.0 0.5 2.2 1.6 1.4
these distributions can be calculated from a, the n u m b e r of organisms originally present, and k, the n u m b e r of species observed. M is the s u m of all the species in all the vessels from an experiment; e.g. M~ is 2+3+0+5+4+5+1+0+6+1=26, M, is 52 and M~ is 13. If contaminants are observed the count is designated b. Then
Lbi,~(v, V)=[ ~hN l[ n2N l( n:~N I (9.5)
XplM,(I_pI),,~N
Mrp2M2( 1 . p2),~N . . M~. ×p3M~(1
166
P3),,~v~-M~ ~"
where N is the number of species originally present, p, is the probability that a particular species will appear in the first set of experiments, ( b l is
a!/b!(a - b)! and Pl = 1 - exp
P2 = 1 - e x p
(10 + b)V ] ) N
(9.6)
(-40Nb)V /
(9.7)
P3 = 1 - e x p / - ~ )
(9.8)
The number of inadvertently added organisms b from the controls is assumed to be the same in the control series as in the experiments. A computer program, available from the authors with instructions, produces an estimate of this value b, together with its standard error, for the experiment sets. At least two sets are required that have different values of a; 10, 40, and 0 in this case. The resulting calculated viabilities V, also given by the program, depend on the estimated number of species N if N is m u c h less than 50. When N is large the calculated viability in the result above is 8.17%, SE = 1.87% with b accounting for 19.6 (SE = 9.0) of the 78 species recognized. This is a sum that m a y include m a n y species that are c o m m o n among the various vessels but different from each other in any particular vessel. The resulting estimate of viability is larger than that from observing growth (Button et al., 1993). An estimate of the total number of species N is required and the calculated viability is slightly different if a smaller N is assumed. For example, in another recent determination from Resurrection Bay, Alaska, three sets of 12 vessels were inoculated with 3 and 12 organisms each along with a control set of 0 organisms inoculated, in the first set 3 vessels were positive with 2, 2, and 1 species present. In the second set, 2 vessels were positive with 1 and 2 species, and the controls remained population free. The calculated viabilities were between 3% and 4% (Table 9.3) and increased only slightly w h e n a smaller number of species was assumed, in which case the calculated error in viability increased as well. Table 9.3 Effect of number of species on the determination of viability N
Viability (%)
Standard error (V)
10
3.91
2.33
25
3.84
1.70
50
3.82
1.63
o~
3.79
1.50
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The population by epifluorescence microscopy was overestimated according to analysis by flow cytometry, perhaps because the fluorescence of the organisms was quite dim in the late fall Alaskan seawater used. Since the assumption of initial population was apparently high and the viability was low, the standard error was large and the calculated viability had a large standard error but the value was not seriously impacted by the number of species assumed to initially be present.
Preparation of dilution medium Good technique and microbiologically impenetrable use of barriers that avoid passage of all bacteria containment is important because a few exogenous contaminants can seriously distort results. Seawater populations are about a million times larger than those initially present in the isolation vessels and minor errors are hard to detect. Such errors inflate apparent viabilities. 1. Wash and further clean by heating to 450 ~, using aluminum foil wrap for small items, two 8 1 carboys, one hundred 50 ml screw top culture tubes, two 47 mm fritted glass filtration systems, siphoning apparatus (including glass tubing, connectors and stopcocks, all with ball joints), filling bells that will fit over a 15 mm filling port in a receiving carboy with ball joints and another to invert over culture tubes when filling, 47 mm glass fiber filters in a Petri dish, and small glass vials for diluting the water sample. Autoclave the following supplies: glass culture tubes, glass vials for dilution, forceps, micropipets, and two 5 x 30 cm strips of non-absorbent cotton. Culture tube caps are lined with Teflon and acid washed. 2. Prepare growth medium for the extinction cultures and for initial dilutions of seawater to the appropriate inoculum size. The procedure involves filtration of seawater to remove large organisms, sterilization by autoclaving, filtration to remove precipitated salts, and transfer to culture tubes. Fit carboy with a rubber stopper, filter holder, glass fiber filter, and bacteriological filter vent. Filter seawater, capped loosely with a glass-tubing-vented rubber stopper wrapped in clean Teflon sheeting, autoclave, and cool over night. Fit carboy with a rubber stopper accommodating an air filter and a port slit and wedged open for a siphon tube. Insert a sterile, glass, siphon apparatus into the carboy through the prepared opening. Close the opening with a hose clamp secured around the stopper. Attach a glass stopcock to the end of the siphon tube to control the flow of seawater during filtration. Wrap all ball joints and protect the end of the siphon tube with sterile cotton to prevent contamination of the medium. Equip the second carboy with a glass filtration assembly and bacteriological air filter to isolate the vacuum pump from the medium. Siphon autoclaved seawater into the second carboy through a 0.2 1.lm 47 mm polycarbonate filter using the sterile cotton and aluminum foil to protect all openings from contamination. A hand pump is used to initiate the siphon. For large volumes of seawater, several filters may be needed.
168
After filtration, aseptically remove the filtration apparatus from the second carboy and insert a sterile siphon assembly terminated with a glass stopcock and filling bell for transfer of medium to culture tubes. 3. Transfer filtered medium to culture vessels. As above, transfer is by siphoning. Care is taken to keep the exposed medium protected from contamination using sterile cotton and aluminum foil. The culture vessels are filled to 40 ml, capped tightly for transport, but loosely for use, inoculated with the desired number of organisms, and incubated at the desired temperature. To reduce chemical contamination by vapor phase transfer during autoclaving microwave sterilization (Keller et al., 1988) may be used. If raw seawater is used, a few bacterial corpses remain visible but most may be distinguished from growing cultures by flow cytometry, and if populations of 10~ml ' or more result they may be detected. However, dissolved organics are added from indigenous bacteria by the high-temperature treatment. These can be eliminated by pre-filtration, but that process can add organics as well, perhaps from chemicals associated with the filters, and damaged organisms. Often populations in inoculated filtered water are observed to be in excess of the original populations present. Using autoclaved medium, the procedure will generate cultures from about 3% of the population although others have indicated much higher success rates (unpublished). Our success rate using microwaved seawater is about the same as with autoclaved seawater.
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Extinction culture in diffusion chambers (Figure 9. I) Dilution bottles may be partitioned against seawater along with its normal complement of microflora for nutrient supply and waste removal.
0 0 ¢0
o
1. Drill Coming Pyrex laboratory bottles (#1395) to give a 1 cm hole in the bottom to accept the inoculum (Figure 9.1). Drill the supplied cap to 25 mm to expose a filter that can retain all bacteria. The inoculating ports are closed with a sterile gauze-wrapped cotton plug. 2. Fit each bottle with a 0.08 pm 47 mm polyester (Poretics) filter, pouring ring, three Teflon gaskets, and the drilled screw-threaded top. The pouring ring, warmed in distilled water for pliability, is fitted over the filter to secure the filter on tile bottle rim. Gaskets, cut from 0.3 mm Teflon sheet with a die, allow the cap to be screwed tightly onto the bottle without disturbing the filter. The bottles are inverted into halfpint jars and autoclaved. Several hours before inoculation, about 20 ml of sterile medium (as prepared above) is added to the half-pint jars and allowed to enter the diffusion chambers through the filter. 3. Determine the bacterial population by epifluorescence microscopy, and prepare the inoculum by diluting the sample to the desired number of organisms with sterile medium. Inoculate through the plugged port of the diffusion chambers with the selected number of organisms (~5 to 50), and suspend the bottles in one-quart canning jars filled with seawater. Rubber bands around the bottle provide a friction
169
"qi ~
-.....
Cotton plug
...... Inoculating
port ~-
~
f-~
<-
a-- Rubber band Culture vessel --- with filtered .... " ~ seawater
<- --1 Pouring ring
~ ~.
<~- ITeflongaskets
~--.---,-"~"
~Raw
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Figure 9.1 Diffusion vessel for extinction culture isolation and culturability determination.
fit to the canning jars for height adjustment to keep the volume of m e d i u m constant. Incubation is on a rocking platform in a softly lighted incubator.
Location of populations Inoculations resulting in growth are located by flow cytometry whereby as few as 10:~ml ~can be determined. Epifluorescence microscopy can be used as well. To estimate the number of species appearing, the number of subpopulation clusters is counted in bivariate histograms of forward scatter and DAP1-DNA fluorescence intensity by flow cytometry. Paired clusters differing in dry mass and DNA content by a factor of two are taken as ln- and 2n-chromosome organisms of the same species. Workers in Giovannoni's lab (personal communication) use molecular probes (Fuhrman and Davis, 1997) to locate populations collected on a membrane clamped below numerous small culture chambers drilled into a Lucite block. Alternatively the number of species might be recognized from qualitative differences in the oligonucleotides produced following restriction and PCR amplification.
Culture storage Due to low nutritional requirements and slow growth, oligobacteria often remain viable in liquid media or on agar plates for m a n y months.
170
Occasional trausfer from liquid media is the safest method of preservation. We retain isolates frozen in glycerol at -55 e, although resuscitation can sometimes take several attempts. The procedure (Gherna, 1994) involves suspending freshly grown cells in glycerol to 20~7~ glycerol, and quickly freezing 1 ml quantities.
Resuscitation Because viability is sometimes low, a whole vial is used. In the case of
Cycloclasticus oligotrophus, the addition of a little substrate speeds the process. Acetate can be used since the organism's specificity for acetate is inherently low; toluene vapors or sterile biphenyl crystals can also be used, alone or in combination with acetate. The thawed contents of the storage vials are dispersed in the appropriate liquid media, usually 50 ml in a conical flask, and the culture incubated until turbid growth appears, usually after one or two weeks.
PROPERTIES OF E X T I N C T I O N AND FUTURE DIRECTIONS
CULTURES om I.
Extinction cultures produce organisms that are smaller in size, lower in solids content and lower in DNA content than enrichment cultures. In situ populations tend to be still smaller in cell size and DNA content. However, specific affinities of cold water organisms for added substrates can be increased 10- to 30-fold by warming for a few hours, during which the cells increase in size. The large-cell-size fraction of the indigenous population is particularly affected.
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CONCLUSION Procedures outlined here are seen as a significant improvement over conventional techniques for isolating and thus understanding pelagic aquatic bacteria. However, much additional improvement, based on systematic experimentation, is required to discuss pelagic aquatic bacteria in more than the most rudimentary detail. The idea that some marine bacteria exist as somni cells or organisms awaiting better conditions appears to have merit because of the vast increase in activity following sampling, particularly in cold waters that are warmed. This activation may be a population-dependent phenomenon particularly associated with waters having a significant annual change in temperature such as high and temperate latitudes. We found large activation energies for amino acid uptake in tile fall and even higher values in the spring (preliminary data). However, most measurements of this phenomenon by others are more modest, giving values equivalent to a normal Q,, for biological systems of near 2. The act of sampling and incubation has a significant effect on the population, which is yet to be understood. Turbulence 171
change is a possible contributing factor to the low viability as well as u n i n t e n d e d chemical changes. That major populations of d e a d cells persist is not well s u p p o r t e d b y a u t o r a d i o g r a p h y observations, by changes in cell properties following s a m p l i n g (in preparation) or b y m e a s u r e d rates of grazing. Thus cultivation of representatives of the n u m e r o u s small l o w - D N A oligobacteria a p p e a r s to be a potentially achievable goal.
References Andrews, P. and Williams. P. J. LeB (1971). Heterotrophic utilization of dissolved organic compounds in the sea. III. Measurement of the oxidation rates and concentrations of glucose and amino acids in sea water. J. Mar. Biol. Assoc. UK 51, 111-125. Britshgi, T. B. and Giovannoni, S.J. (1991). Phylogenetic analysis of natural marine bacterioplankton population by rRNA cloning and sequencing. Appl. Environ. Microbiol. 57, 1701-1713. Button, D. K. (1998). Nutrient uptake by microorganisms according to kinetic parameters from theory as related to cytoarchitecture. Microbiol. Mol Biol. Rev. 62, 636-645. Button, D. K. and Robertson B. R. (1993). Use of high-resolution flow cytometry to determine the activity and distribution of aquatic bacteria. In: Handbook of Methods in Aquatic Microbial Ecology (P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 163-173. Lewis, Ann Arbor, MI. Button, D. K. and Robertson, B. R. (2000) Effect of nutrient kinetics and cytoarchitecture on bacterioplankter size. Limnol. Oceanogr. 45, 499-505. Button, D. K., Schut, E, Quang, P., Martin, R. M. and Robertson, B. (1993). Viability and isolation of typical marine oligobacteria by dilution culture: Theory, procedures and initial results. Appl. Environ. Microbiol. 59, 881-891. Cammack, R. (1997). The enzyme at the end of the food chain. Nature 390, 443-444. Daley, R. l- and Hobbie, J. E. (1975). Direct counts of aquatic bacteria by a modified epi-fluorescent technique. Limuol. Oceanogr. 20, 875-882. Fuhrman, J. A. and Davis, A. A. (1997). Widespread Archaea and novel bacteria from the deep sea as shown by 16S rRNA gene sequences. Mar. Ecol. Prog. Set. 150, 275-285. Gasol, J. M., Zwiefel, U. L., Peters, E, Fuhrman, J. A. and Hagstr6m, A. (1999). Significance of size and nucleic acid content heterogeneity as measured by flow cytometry in natural planktonic bacteria. Appl. Environ. Microbiol. 65, 4475-4483. Gherna, R. L. (1994). Culture preservation. In: Methods for General and Molecular Bacterioloy,y (P. Gerhardt, R. G. E. Murray, W. A. Wood, N. R. Krieg, Eds). American Society for Microbiology, Washington, DC. Jannasch, H. W. (1979). Microbial ecology of aquatic low nutrient habitats. In: Stratc\e,ies of Microbial Life in Extreme Environments (M. Shilo, Ed.). Dahlem Konferenzen, Berlin. Keller, M. D., Bellows, W. K. and Guillard, R. R. L. (1988). Microwave treatment for sterilization of phytoplankton culture media. J. Exp. Mar. Biol. Ecol. 117, 279-284. Quang, P., Button, D. K. and Robertson, B. R. (1998). Use of species distribution data in the determination of bacterial viability by extinction culture of aquatic bacteria. J. Microbiol. Methods. 33, 203-210. Rice, J. A. (1995). Mathematical Statistics amt Data Analysis, 2nd edn. Duxbury Press, Belmont.
172
Robertson, B. R., Arhelger, S., Kinney, P. J. and Button, D. K. (1973). Hydrocarbon biodegradation in Alaskan waters. In: The Microbial Degradation of Oil Pollutants. Center for Wetland Resources (D. G. Ahearn and S. P. Meyers, Eds). Baton Rouge, LA. Robertson, B. R., Button, D. K. and Koch, A. L. (1998). Determination of the biomasses of small bacteria at low concentration in a mixture of species with forward light scatter measurements by flow cytometry. Appl. Environ. Microbiol. 64, 3900-3909. Schmidt, T. M., DeLong, E. E and Pace, N. R. (1991). Analysis of a marine picoplankton community by 16S rRNA gene cloning and sequencing. [. Bacteriol. 173, 4371-4378. Sieracki, M. E. and Viles, C. L. (1992). Distributions and fluorochrome-staining properties of submicrometer particles and bacteria in the North Atlantic. DeepSea Res. 39, 1919-1922. Torrella, E and Morita, R. Y. (1981). Microcultural study of bacterial size changes and microcolony and ultramicrocolony formation by heterotrophic bacteria in seawater. Appl. Environ. Microbiol. 41, 518-527. Whitesides, M. D. and Oliver, J. D. (1997). Resuscitation of a Vibrio wdnificus from the viable but nonculturable state. Appl. Environ. Microbiol. 63, 1002-1005. Zobell, C. E. and Upham, H. C. (1944). A list of marine bacteria including descriptions of sixty new species. Scripps Institution of Oceanography Contributions 5, 239-292. om
List of suppliers
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173
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10 Determining the Physiological Status of Individual Bacterial Cells H H Howard-Jones, HE Frischer and PGVerity Skidaway Institute of Oceanography, I 0 Ocean Science Circle, Savannah, Georgia 3141 I, USA eeeeeeeeeeeeeeeeeeeeeeeeeeeeeeoeoeeeeeoeoeeeoeoe
CONTENTS Preface General introduction Vital stain and probe (VSP) method Nucleoid staining technique Microautoradiography and probes Live/dead BacLight TM BacteriaIViability Kit Direct viable counting: Kogure method Conclusions
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PREFACE O
During the past several decades the field of marine microbiology has blossomed, arguably due to the development and application of new methodological approaches. During this period our awareness of the importance, diversity, and complexity of microbial communities in marine systems has dramatically increased. This awareness has prompted the recognition that methods capable of assessing the identity and activity of individual cells are needed to augment those available to measure bulk community properties. Thus, an active research area has been the development of new methods to measure the physiological status and activity of individual cells in planktonic environments. Already application of these methods has led to potentially paradigm shifting observations regarding the composition and activity of marine microbial assemblages. For example, it is now reasonably well accepted that a significant fraction of planktonic bacteria at any given instance in space and time are either dead or relatively inactive. In this chapter we describe and provide detailed laboratory protocols for five techniques that can be used to estimate the physiological status of individual cells in marine and estuarine water samples. M E T H O D S IN MICROBIOLOGY, V O L U M E 30 ISBN 0 12-521530 4
C o p y r i g h t © 2001 A c a d e m i c Press Ltd All rights of reproduction m a n y form reserved
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eeeeee, GENERAL INTRODUCTION During the past quarter century, the appreciation of bacteria as important biological components in marine ecosystems has undergone a dramatic revolution. This epiphany has largely been stimulated by two methodological advancements; direct microscopic enumeration techniques to estimate bacterial abundance (Hobbie et al., 1977; for a recent review see Kepner and Pratt, 1994) and radiotracer methods for estimating the activity of bacterial communities (Brock and Brock, 1966; Carman, 1990). The application of these approaches to marine sciences has led to a paradigm shift regarding the importance of microbial communities in marine systems. For example, prior to the development of these techniques bacteria were thought to be relatively unimportant in marine systems. We now recognize that a majority of living biomass in the sea consists of bacteria and, on average, half of all photosynthetically fixed carbon is processed through heterotrophic bacteria and the microbial loop (Whitman et al., 1998; Ducklow, 1999). However, as the awareness of the importance, diversity, and complexity of microbial communities has developed, it is now recognized that an understanding of the ecology of microbial communities in nature requires information about the identity and activity of individual cells (Sherr et al., 2000). Such information would allow us to address questions such as: 'Which cells are active, why, when, and what are the controls?' Traditional counting and activity measurements cannot provide answers to these and similar questions; therefore, a considerable amount of recent research effort has focused on the development of new techniques that permit estimation of the activity of individual bacteria cells in marine samples. We predict that, just as direct enumeration and activity measurements transformed the field of marine microbiology in the last several decades, the capability of measuring the activity of individual bacteria cells will likewise propel the field to new levels of understanding. Thus, the object of this chapter is to describe several methodologies that are currently under development a n d / o r in use to investigate the physiological activity of individual bacteria cells in marine samples. Several cellular criteria have been proposed and targeted by specialized stains and probes as indicators of cellular activity. These vital stains and probes take advantage of a number of different cellular properties that can be associated with cellular physiology. Examples of criteria that have been used to assess the metabolic status of individual cells include membrane integrity (L6pez-Amor6s et al., 1995; Mason et al., 1995; Haugland, 1996; L6pez-Amor6s et al., 1997), cellular reducing potential (Zimmermann et al., 1978; Rodriguez et al., 1992; Epstein and Rossel, 1995), specific enzyme activity (Lundgren, 1981; Tsuji et al., 1995), and ribosomal RNA (rRNA) cell content (Schaechter et al., 1958; Bremer and Dennis, 1987; DeLong et al., 1989; Lee et al., 1993; Williams et al., 1998). Although several techniques are available and have been used to describe the activity of individual cells in a variety of environments, currently there is no clear consensus among the scientific community regarding the absolute validity of these methods, or even which methods 176
are the most reliable (but see Karner and Fuhrman, 1997). Therefore, at this time, regardless of the method used, it must be recognized that the physiological state of a cell is operationally defined and inferred from the general properties (specificity, sensitivity, etc.) of a particular method, and that the results produced from any single method may provide only relative rather than absolute information. Despite the recognized limitation in the methods available for determining individual cell metabolic status, a number of studies have examined the activity of cells in a variety of marine environments. A general review of this literature suggests that regardless of the method employed, it is clear that a significant fraction of cells visualized by direct epifluorescence microscopy methods are probably dead or relatively inactive (del Giorgio and Scarborough, 1995; Zweifel and Hagstrom, 1995; Heissenberger et al., 1996; Choi et al., 1999; Sherr et al., 1999; Sherr et al., 2000). However, the relative proportion of these groups of cells remains a subject of active debate. Estimates of the proportion of dead and low-activity cells range from 0.1 to 99% depending on which method was used and what environments were examined (Lovejoy et al., 1996; Karner and Fuhrman, 1997; Sherr et al., 2000). These results confirm the hypothesis that techniques that measure bulk community properties underestimate the complexity of microbial community structure and function, and that methods to assess the physiological status of individual cells in situ are required. The purpose of this chapter is to provide a summary and detailed protocols for several methods that show promise for measuring the activity of individual cells in marine samples. We have not discussed the use of CTC within this chapter since this method is described in an earlier chapter of this book (Sherr et al., Chapter 8, this volume). It is our hope that as these methods are improved, calibrated, and commonly used, a more consistent and coherent understanding of the activities of individual bacterial cells in situ will be elucidated.
• ~ , ~
V I T A L S T A I N A N D PROBE (VSP) M E T H O D
Introduction The Vital Stain and Probe (VSP) method is an epifluorescent direct microscopy counting technique that combines the general DNA stain DAPI, the vital stain propidium iodide (PI), and 16S rRNA targeted oligonucleotide probes to identify bacterial cells in situ (Williams et al., 1998). This technique simultaneously assesses two independent criteria of cells that can be related to cell physiological status: (a) membrane integrity and (b) cellular ribosomal content. Membrane integrity is assessed using the vital stain PI. PI is a nucleic acid stain that accumulates in cells that have leaky or compromised membranes (Jernaes and Steen, 1994; Lopez-Amoros et al., 1995, 1997; Williams et al., 1998). Cells that stain positively with P1 are interpreted as being dead, although recently divided cells may also be stained by PI (Howard-Jones et al., unpublished). The activity status of microbial cells is determined by hybridiza177
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tion with 16S rRNA oligonucleotide-targeted probes (Lane et al., 1985; Braun-Howland et al., 1992; Spring et al., 1993; Amann et al., 1995). Cells that contain sufficient numbers of ribosomes to be detected by probe hybridization can be visualized with epifluorescent microscopy. Those that appear positively hybridized with oligonucleotide probes are considered to be active since activity and growth rate can be qualitatively related to ribosome content (Giovannoni et al., 1988; DeLong, 1989; Lee et al., 1993, 1994). The combination of DAPI, PI, and probes reveals the physiological status of individual cells. Thus, it is hypothesized that the VSP method can distinguish between four categories of cells: live and active, dead, dead but recently active, and cells that are living but relatively inactive (Table 10.1). Since two independent cellular criteria are simultaneously evaluated by the VSP method, this technique is considered fairly robust in comparison to methods that rely on single cellular criteria. Similar characterization of cells has been defined by Gasol et al. (1999) as follows: (1) large, rapidly growing cells, or cells defined as live and active by the VSP method, (2) dead cells with no potential for growth correspond to those characterized as dead by the VSP; and (3) cells that are growing at a very low rate, or cells that are inactive possibly due to inappropriate growth conditions defined as two separate groups by Gasol et al. (1999) correspond to cells that are live but express low activity as defined by VSP technology. Gasol et al. (1999) also defines a group of cells as cell fragments or ghost cells. This group of cells would be characterized by two different categories of the VSP: (a) cell fragments, if they contain DNA would be classified as dead, while (b) ghost ceils would be categorized as living but low activity cells.
Table I 0. I Proposed definitions of cell status as determined by the uptake or exclusion of propidium iodide (PI) and hybridization with 16S rRNA targeted oligonucleotide probes Cell status
DAPI
PI
Probe
Live, active Dead Dead, recently active Live, low-no activity
+ + + +
+ +
+ +
The VSP method uses 16S rRNA targeted oligonucleotide probes which are comprised of synthesized DNA oligomers, a fluorochrome, and a linker molecule. Oligonucleotides are custom-synthesized with a 5' aminoqink modifier and conjugated with a fluorochrome in the laboratory. Alternatively, fluor conjugated oligonucleotides can be synthesized directly. Oligonucleotide probes used in our laboratory are synthesized by Integrated DNA Technologies (Coralville, IA) and were conjugated with fluorescein isothiocyanate (FITC). However, a variety of other fluorochromes are available, including FluorX, the cyanine dyes (Cy3, Cy5), 178
tetramethvl-rhodamine isothiocyanate (TRITC) etc. (Braun-Howland et a/., 1992; Hicks et al., 1992; DeLong, 1993; Amann et al., 1995; Glockner et al., 1996). A wide range of phylogenetically-specific 16S rRNA targeted oligonucleotide probe sequences have been described in the literature that are suitable for use with the VSP method. Therefore, the VSP technique has the potential to measure the activity and viability of individual cells within the bacterial population, the activity of individual subgroups, or the activity of the entire community, depending on the specificity of the probes utili×ed.
Principle DAPI is a general fluorescent stain that complexes to A-T rich sequences in the major groove of dsDNA in bacterial cells. Under UV (Exciter filter BP 330-385, dichroic mirror DM 400, and barrier filter BA 420) excitation, DAPI stains the DNA of bacteria cells white-blue (emission wavelength for white blue is @ or above 390 nm) and differentially stains particulate matter (detritus) yellow (emission wavelength for yellow is below 390 nm). The vital stain propidium iodide (PI) binds to nucleic acids in cells that have lost membrane integrity (Jernaes and Steen, 1994; LopezAmoros cta/., 1995, 1997; Comas and Vives-Rego, 1997; Williams et aI., 1998). PI accumulates in cells that have compromised membranes and stains the cell a red color when visualized with Wide Green (WG~,) Excitation filter 510-550, dichroic mirror DM570, and barrier filter BA 590) excitation. Cells that are visualized in the Pl field are considered dead. Propidium iodide passively diffuses into cells (influx), a process driven by the dye concentration across tire cell wall. However, in cells with intact membranes this process is limited and possibly counter-balanced by an efflux system [driven by the electrochemical potential] in cells that are considered live by definition of an undamaged membrane (Jernaes and Steen, 1994). Active cells do not stain with PI (Lopez-Arnoros et al., 1995). The ribosomal RNA content of the cell is evaluated by employing 16S oligonucleotide targeted probes. Cells that contain sufficient RNA to be considered metabolically active are visualized green with Narrow Blue (NBX) Excitation filter BP 470-490, dichroic mirror DM 500, and barrier filter BA 515) excitation. The following method yields quantitative information regarding the viability and physiological status of individual bacterial cells iH sitzl.
Equipment and reagents Vacuum manifold, 25 m m vacuum filter holders. Epifluorescence microscope equipped w i t h a UV, Wide Green (WG), and Narrow Blue (NB) filter sets. • Vacuum pump (source of vacuum). • I n c u b a c o r able co maintain 37°C temperature. • •
•
Coplin slide staining jars.
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1.5 and 4 ml Eppendorfcubes.
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•
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•
•
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Humidified chamber (covered tupperware dish with paper towels lining the bottom; soaked with distilled water). 0.2 ~tm black polycarbonate filter (Micron Separation, Inc.). 25 mm GF/F (Whatman). 99% glycerin. 100% analysis grade methanol. 10 mM MgSO4 (pH 6.5); this solution can be pre-made and stored at room temperature; prior to use, 10 mM MgSO4 is filtered through a 0.2 lum Acrodisc filter. Ix SET (I 50 mM NaCI, 20 mM Tris-HCI, I mM EDTA, pH = 8.0); this solution is pre-made and stored in an incubator at 37°C; Ix SET is filtered through a 0.2 lure Acrodisc filter. Ethanol: Formaldehyde (90:10) (100% EtOH, 37% formalin). Hybridization solution (contains the following: 0.2% w/v bovine serum albumin (BSA), 0.01% w/v polyadenylic acid, Ix SET, I1% dextran sulfate); hybridization solution can be pre-made and stored in individual I ml aliquots in the freezer at -20°C. DAPI (stock solution: 50 lug ml ');the stock solution is made by dissolving 5 mg of DAPI in 100 ml of distilled water; the solution is filtered through a 0.2 lum Acrodisc filter and stored in the dark at 4°C. Propidium iodide, PI (stock solution: 20 lug ml ');the stock solution is made by dissolving I mg of PI in 50 ml of distilled water; the solution is filtered through a 0.2 lum Acrodisc filter and stored in the dark at 4°C. Fluorochrome labeled oligonucleotide probes; this method employs three distinctly different oligonucleotides targeted to universally conserved regions of the bacterial ribosome. Only three oligonucleotides are used for this method, however, there are numerous sequences available for synthesizing 16S rRNA probes (Lane et al., 1985; Giovannoni et al., 1988; Lee et al., 1993; Amann et al., 1995; Frischer et al., 1996). Each of these is conjugated to the fluorochrome FITC (fluorescein isothiocyanate): (a) Primer A (5'-gwattaccgcggckgctg; Lane et al., 1985) 5 1 9 - 5 3 6 (b) Primer C (5'-acgggcggtgtgtrc; Lane et al., 1985) 1 3 9 2 - 1 4 0 6 (c) EUB 342 (5'-ctgctgcsycccgtag;Vescio and Nierzwicki-Bauer 1995) 3 4 2 - 3 5 8 The numbers in italics correspond to the nucleotide positions relative to the E. coli 16S rRNA gene.Ambiguous bases are w = a or u; r = a or g; s = c or g; k = g or u; y = c or u.The three oligonucleotide probes are combined (three probes per 1.5 ml Eppendorf tube; 340 ng each), dried in a vacuum dessicator and stored in the freezer at -20°C. Glass microscope slides. Glass coverslips. Microscope grade non-fluorescence immersion oil.
Protocol 1. Water samples (15ml) are collected in 20ml scintillation vials containing 5 ml of sterile glycerol (99%) for a final concentration of 25%; vortex well; allow the sample to equilibrate at room temperature for approximately 10 min before analysis or storage at -20°C. Water samples in 25% glycerol can be stored for 30-60 days at -20°C. 180
2. If necessary (culture work), cells are diluted appropriately in 10 mM MgSO~ (pH 6.5). 3. For sufficient statistical treatment, triplicate samples are prepared and enumerated. 4. Cells are fixed with analysis grade methanol (final concentration: 2~:). Methanol fixation stabilizes cell membranes so that cells remain intact throughout the protocol treatments. 5. Incubate cells for 30 min at room temperature, preferentially in the dark. 6. Stain cells simultaneously with 60 btlml' of sample DAPI (final concentration: 3 btg ml ~) and 10 btl ml ~PI (final concentration: 0.2 btg ml '). Stain samples in the dark for 30 min at room temperature. 7. Permeabilize the cells with ethanol: formaldehyde (90:10) (100 sample original volume). Oligonucleotide probes are 'large' molecules that need to enter the cell in order to hybridize with the ribosomal subunits. Permeabilization is required because it serves to 'poke holes' into the membrane, allowing the probe molecules to enter into the cell more efficiently and easily. 8. Incubate in the dark for 30 rain at room temperature. 9. Filter the samples onto a 0.2 btm black polycarbonate filter underlaid with a 25ram G F / F filter. Filter the samples under low vacuum (<15 m m Hg). The underlying G F / F filter helps to produce an even distribution of cells on the polycarbonate filter. 10. Wash the filters three times with 1 ml of 10mM MgSO~ [pH 6.5] to remove any non-specifically bound stains (under low vacuum < 15 m m Hg). 11. Place the filter onto a labeled glass microscope slide, cell side face up. Attach the polycarbonate filter to the glass slide with small drops of fingernail polish at the edges of the filter. 12. Place the slides into a humidified chamber and overlay the filters with 500 btl of the hybridization/probe solution. The hybridization/probe solution is a combination of the hybridization solution and the three oligonucleotide probes. When it is time to hybridize, one aliquot of hybridization solution is added to one probe (consists of three oligonucleotide, 340ng each) and vortexed well. For each 1 ml mixture, two slides can be overlaid with hybridization/probe solution. 13. Place chamber into an incubator equilibrated to the appropriate hybridization temperature (for the probe set described here, 37°C) and incubate tile slides overnight. A m i n i m u m of 8 h is required for hybridization. 14. Next day: remove the slides from the chamber and wash them for 30 min in three changes of lx SET (in a Coplin jar at the hybridization temperature). 15. Lay slides out to air dry (5 rain) at room temperature. 16. Place a small drop of mounting solution (50:50 lx SET, glycerol) on the filter and cover with a glass coverslip. 17. Enumerate cells with an epifluorescent microscope equipped with the following filter sets: 181
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(a) DAPI: Wide UV (WU). Olympus U-M536 contains exciter filter BP 330-385, dichroic mirror DM 400, and barrier filter BA 420. (b) PI: Wide Green (WG). Olympus U-M526 contains exciter filter BP 510-550, dichroic mirror DM 570, and barrier filter BA 590. (c) Probe (FITC or FluorX): Narrow Blue (NB). Olympus U-M536 contains exciter filter BP 470-490, dichroic mirror DM 500, and barrier filter BA 515. Enumeration is facilitated by using an imaging system equipped with a 100x Plan-apochromat oil objective (NA 1.3-1.4), a color integrating analog or digital CCD, and commercial (e.g. Image Pro Plus) or custom (Sieracki et al., 1989, Shopov et al., 2000) imaging software. Systematic evaluation of staining protocols during the development of the VSP method demonstrated that removing the P! stain (washing step) from the sample incubation, prior to permeabilizing the membranes for probe hybridization, did not influence the fraction of cells appearing dead or dead but recently active (Howard-Jones et al., unpublished). Therefore the protocol does not require removal of PI by washing. Because the fluoresence emission spectra of PI overlaps that of chlorophyll a, autofluorescent picoplankton may be identified as dead cells. However, in our experience, bacterial cell abundance is usually one to two orders of magnitude greater than the abundance of autofluorescent picoplankton cells in marine water samples. Therefore, confusion of PIstained bacteria and autofluorescent picoplankton is rarely a problem. Nevertheless environmental samples that have fewer bacterial cells, requiring larger volumes of water to be filtered for sufficient numbers of cells, might pose a problem. It is recommended that the number of autofluorescent picoplankton be determined in each sample prior to VSP analysis. An example of VSP staining of bacterioplankton from surface waters from the Barents Sea is shown in Plate 2. The micrographs represent cells that are stained with DAPI (left), propidium iodide (right), and universally-targeted 16S rRNA oligonucleotide probes conjugated to FITC (centre). Cells that stain with both DAPI and P! are considered dead while cells that appear in both the DAPI and probe fields are considered active. Cells that appear in both the probe and PI fields are cells that are categorized as dead but recently active, or cells that are undergoing division or recently divided. Cells that fall into the category of dead but recently active are those that have in all probability been active within a few days of sampling. These cells contain a sufficient amount of RNA to be detected with probes, however, their cell membranes have been significantly compromised. Cells that are undergoing division or have recently divided can stain positively for both PI and probe. Based on ongoing laboratory and field studies, recently divided cells persist only for minutes, while recently active cells are present for hours to days (Hong et al., 1998; Howard-Jones et al., unpublished). Advantages Method measures two independent criteria simultaneously. Individual cells are scored as active, inactive, dead or live.
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Accurately and reliably distinguishes between living and dead cells in manipulated and natural systems. • Samples can be collected and stored for up to 3 months. • Can be used to assess the activity of the total bacterioplankton community or specific fractions of the community depending on the probes used.
•
Disadvantages Under some circumstances propidium iodide can yield variable staining of cells.Therefore, the scoring of Pl staining may require a subjective decision. Only cells that appear brightly stained with PI are considered dead. Use of a computer-aided image analysis system can minimize this difficulty. • Except in a few special cases, ribosome-targeted probes rarely provide functional information about the individual cells. • The relationship between ribosomal content and activity varies among cell types. •
.e~l, e e e N U C L E O I D S T A I N I N G T E C H N I Q U E Introduction In a healthy bacterial cell DNA is organized in a condensed structure called a nucleoid. The nucleoid can be observed with epifluorescence microscopy w h e n it is stained with a D N A fluorochrome such as DAPI. However, because DAPI is a general stain that can bind non-specifically to bacterial cells and debris1 counts using this technique are thought to overestimate the n u m b e r of cells that actually contain DNA (Zweifel and Hagstrom, 1995; Choiet al., 1996; Vosian and van Noort, 1998). The goal of enumerating nucleoid-containing cells (NUCC) is to count only the cells that contain visible DNA and exclude bacterium-like particles or dead cells ('ghosts') that might be counted after staining with standard DAPI m e t h o d s (Karner and Fuhrman, 1997). Cells that contain nucleoids are considered active, although NUCC bacterium counts probably represent m a x i m u m estimates of the n u m b e r of active bacteria since the presence of a nucleoid does not ensure that a bacterium is growing or even capable of growth (Zweifel and Hagstrom, 1995). For example, cells that are dormant, starved, or infected by viruses m a y contain nucleoids although they are likely to be relatively metabolically inactive. Conversely, cells that are inactive but alive m a y be identified as dead cells (Proctor and Fuhrman, 1990; Zweifel and Hagstrom, 1995; C h o i e t al., 1996, 1999). For example, Choi st ~71. (1996) found that starving, non-nucleoid cells maintained in the laboratory regained yellow-fluorescing nucleoids w h e n supplied with a nutrient source, while with natural samples, approximately 20~/~ of the non-nucleoid cells developed visible nucleoids following the addition of nutrients. These results suggest that non-nucleoid cells m a y still be viable but contain D N A in a less compact region of the cell or of insufficient concentration to cause DAPI-stained DNA to precipitate (Choiet al. 1996).
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The technique described here to visualize nucleoid containing cells was developed by Choi et al. (1999) and represents a modification of the method first described by Zweifel and Hagstrom (1995).
Principle DAP1 is a fluorochrome that binds to the A-T rich sequences in dsDNA. However, standard methods of using DAPI to enumerate bacteria are challenged by the demonstration that it binds non-specifically, particularly at salinities greater than 12 ppt. DAPI binding most likely occurs on reactive bacterial surfaces, rather than to the DNA within the cell membrane (Zweifel and Hagstrom, 1995). Staining bacterial cells in fresh water promotes effective binding of DAP! to DNA (Choi et al., 1996). To remove non-specifically bound DAPI, DAPI-stained cells are washed with a solution of 2-propanol. This washing step removes DAPI from a large but variable number of cells (Zweifel and Hagstrom, 1995; Choi et al., 1996; Vosjan and van Noort, 1998). Fixation prior to de-staining the cells is crucial because it provides physical rigidity to the bacterial cell membranes and allows fluorochromes to penetrate the membranes more readily. Following this protocol, nucleoid-containing cells are visualized as bright yellow spots within dimly blue-fluorescing cells (Choi et al., 1996). The following method yields estimates of viability and metabolic activity based on the presence of a condensed region of DNA known as the nucleoid.
Equipment and reagents • Vacuum manifold, 25 mm vacuum filter holders. • Epifluorescent microscope equipped with UV capability. • Temperature-controlled water bath. • 25 mm, 0.2 ~m black polycarbonate filters (Micron Separations, Inc.). • GF/C filters 25 mm (Whatman). • 0.2 lum pore-size filtered Millipore Milli-Q water. • Fresh sterile-filtered 37% borate-buffered formalin. • DAPI (stock solution: 0.1 mg ml-').The stock solution is made by dissolving 5 mg of DAPI in 50 ml of distilled water, filtered through a 0.2 pm filter, and stored in the clark at 4°C. • Pro-analysis grade 2-propanol warmed to 60-65°C. • Glass microscope slides. • Glass coverslips. • Microscope grade immersion oil.
Protocol 1. Collect water samples in 1 1bottles. 2. Separate sample into two aliquots: one that will be stained immediately to determine total cells and one that will be treated as stated 184
below. For the sample that is to be stained immediately, the cells are fixed with a 5% final concentration of 37% formalin, stained with 10 gg ml ~final concentration of DAPI for 10 min and filtered onto a 25 mm, 0.2 g m black polycarbonate filter. The filters are then m o u n t e d on slides u n d e r oil and coverslips, and e n u m e r a t e d by standard UV epifluorescence microscopy. 3. To visualize nucleoids, fix samples in glass containers with 10% final concentration of 37% formalin. Vortex the samples thoroughly and allow them to equilibrate at room temperature for 10 rain. The cells are fixed with formalin to avoid swelling of the cells and to avoid significant cell loss in the staining process. 4. Set u p the filter towers with 0.2 gm black polycarbonate filters underlain with G F / C filters. The G F / C filters help to produce an even distribution of cells on the polycarbonate filter. 5. Diluting the sample: (a) Low salinity samples (< 12 ppt). Pipette sample (1-2 ml of natural water) on to the filter tower. A d d 5 ml of 0.2-gm-filtered MilliQ water. (b) High salinity samples (> 12 ppt). Dilute the sample with 0,2 ~m Milli-Q water by pipetting 5 ml of 0.2-Hm-filtered MilliQ water to the filter tower, followed by the sample (1-2 ml of natural water). Another 5 ml (depending on the bacterial abundance) of filtersterilized water to the tower can be used as an additional rinse. The addition of distilled water is required for (i) lowering the salinity to less than 12 ppt and (ii) removing remaining a l d e h y d e fixative. 6. Filter the samples until dry at 10 m m Hg. 7. Add DAPI (final concentration: 10 gg ml ~ sample) to the filter tower and incubate for 1 0 m i n in the dark (i.e. cover the filter tower completely with a l u m i n u m foil). 8. Filter d o w n the samples to dryness at 10 m m Hg. 9. Apply three, 1 ml rinses of w a r m (60-65°C) 2-propanol to the filter towers. It is important to rinse gently to avoid dislodging cells from the filter. The propanol wash removes the non-specifically bound DAPI stain from the cells resulting in only nucleoids being stained with DAPI. 10. Once the filter is dry, remove and place it on a paper towel to allow excess propanol to evaporate. 11. Place the filter onto a microscope slide. Immersion oil is placed both u n d e r and on top of the filter before it is covered with a glass coverslip. If necessary, samples can be stored at -20°C for u p to several weeks. 12. Cells are e n u m e r a t e d with an epifluorescent microscope equipped with a UV filter set (e.g. Zeiss 4877-02, excitation filter 365 n m / b a r r i e r filter 420 nm). An example of nucleoid visualization of marine bacterioplankton is s h o w n in Figure 10.1.
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Figure 10.1. Nucleoid staining technique. DAPI-stained marine bacteria; total numbers compared with NUCC bacteria as viewed by epifluorescence microscopy. (A) Conventional DAPI stained sample (site NB1). (B) Bright NUCC bacteria in the same sample used in panel A, after de-staining (overexposure creates images that are larger than the original cell). (C) Nucleoids in a marine isolate (ZS2) visualized by superimposing a binary image of bright nucleoids onto the original negative to compensate for the color information that was lost in transforming to gray scale, Total counts of marine bacteria include a large fraction of non-nucleoid-containing bacteria (ghosts). Reprinted with permission from: Zweifel and Hagstrom (1995), Applied Euviroumeut Microbiolo~(ll.
Advantages • •
•
Technique is comparatively simple and rapid. Theoretically provides information on total bacteria and active bacteria, regardless of cell type, although this assumption has not been rigorously tested. Data is more relevant to community dynamics than just total DAPI-stained cell abundance.
Disadvantages t
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Unable to discriminate living from low activity cells. Cells that are living but inactive (e.g. starved cells) can be misidentified as either living/active or dead cells. Cells that do not contain visible nucleoids are difficult to discern. Separate samples are required for determining total cell counts. Method relies on a single criterion.
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MICROAUTORADIOGRAPHYAND
PROBES
Introduction Microautoradiography is a technique that detects the cellular localization of a radiolabeled substance ill situ (Brock and Brock, 1966; Hoppe, 1976; Meyer-Reil, 1978). Specifically, microautoradiography can be used in radiotracer studies to determine the proportion of aquatic microorganisms that are metabolizing a given radiolabeled substrate (Carman, 1990). The labeled substrate should be a building-block molecule used bv the cell only to make the molecules of interest. For example, radiolabeled thvmidine is used as a measure of DNA synthesis, uridine for RNA synthesis, and amino acids for protein synthesis (Wolfe, 1983). Although it has been recognized that these assumptions are idealized and that the radiolabeled compounds can enter into numerous other biosynthetic pathways, for the present discussion, it is assumed that the majority of these molecules go directly into cell synthesis. Microautoradiography relies on the ernission of beta particles ejected during radioactive decay. Beta particles can expose crystals (silver grains) in a photographic emulsion that corresponds spatially to ceils that have incorporated the radiolabeled compound. After incubation and development, the cells can be examined microscopically to distinguish metabolically active from inactive cells (Brock and Madigan, 1988). The method described here combines microautoradiography with 16S ribosomal RNA-targeted oligonucleotide probes which allows simultaneous in situ identification and determination of substrate uptake patterns of individual cells (Lee eta/., 1999; Ouverney and Fuhrman, 1999; Cottrell and Kirchman, in press). Cells are labeled with a general stain (DAPI), fluorescently labeled oligonucleotide probes, and tritiated carbon sources. This technique incorporates two independent measurements: (a) cellular ribosomal content/identity and (b) substrate uptake. The combination of microautoradiography and phylogenetic-specific probes provides a means to identify cells that are actively incorporating a specific substrate. This approach promises to become an extremely valuable tool for determining the functional role of diverse microbial communities in nature.
Principle DAPI is a general DNA stain that binds to the A-T rich sequences in dsDNA and cells that are stained with DAP1 can be visualized by epifluorescence microscopy. DAPI staining provides a means to enumerate the total bacterial community regardless of identitv or physiological status. 16S rRNA oligonucleotide probes bind to ribosomal RNA in phylogenetically specific groups of cells that can be considered to be metabolically active (see VSP section of this chapter). Microautoradiography is used to identify cells that are actively incorporating a specific substrate. The method described here is based primarily on the STARFISH protocol developed by Ouvernev and Fuhrman (1999), although similar methods
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have been developed by other groups that incorporate several modifications and or improvements to this method (Lee et al., 1999; Cottrell and Kirchman, 2000). For example, in the MICRO-FISH method described by Cottrell and Kirchman (2000) a special slide having a viewing hole is not required. As described in their manuscript, Ouverney and Fuhrman (1999) utilized the STARFISH technique to examine the utilization of glucose and free amino acids by eubacteria c~-proteobacteria, and the Cytophaga-Flavobacterium sub-groups of the eubacteria. Cottrell and Kirchman (2000) utilized the MICRO-FISH method to examine the utilization of low and high molecular weight dissolved organic matter (DOM) by eubacteria, the Alpha-, Beta-, and Gamma-subdivisions of the proteobacteria, the Cytophaga-Flavobacter cluster of the CFB division, and the high G+C subdivision of the Gram positive phylum.
Equipment and reagents
• • • • • • • • • • • • • • • • • •
• • •
Refrigerator (+4°C). Freezer (-20°C). Temperature controlled water bath. Epifluorescence/transmission light microscope equipped with UV and Wide Green (WG) filter sets. Rotary shaker. Vacuum manifold, 25 mm vacuum filter holders. Vacuum source. Scintillation counter. Dark room. ITT NightVision scope. 15 W safelight. 250 ml acid washed dark bottles. Sterile conical tubes (minimum 50 ml volume). Micropipettes. Forceps. 25 mm 0.2 ~m Nuclepore polycarbonate filters. 25 mm 0.8 lum typeAA Millipore filters. Heavy Teflon glass slides with ten 7 mm diameter wells. Lightproof box wrapped in aluminum foil. Tritiated glucose, specific activity 50 Ci mmol ', final activity 10 pCi (Dupont, N ET100). Tritiated amino acid mixture, specific activity 50 Ci mmoF', final activity 10 IJCi (Amersham, St Louis, MO, TRK440). DAPI (stock concentration: 0.1 ~g ~1 '). The stock solution is made by dissolving 5 mg of DAPI in 50 ml of distilled water. The stock solution can be stored in the dark at 4°C. 0.2x SET: (I × SET is 150 mM NaCI, 20 mM Tris-HCI pH 7.8, and I mP1 EDTA) I × PBS (phosphate-buffered saline: 8 g NaCI, 0.2_g KCI, 1.44 g Na2HPO 4,0.24 g KH2PO4, pH = 7.4). Oligonucleotide probes labeled with the fluorochromeTRITC (5 ng ml ~each of); probes can be labeled with a variety of other available fluorochromes such as Cy3:
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(a) Negative control [5'-cctagtgacgccgtcgac] (minimum of three mismatches with all rRNA sequences in the Ribosome Database Project) (b) Universal [5'-gwattaccgcggckgctg] 519-536 (c) Bacteria [5'-accgcttgtgcgggccc] 342-359 (d) (z-proteobacteria [5'-cgttcgytctgagccag] 19-35 (e) Cytophaga-Flavobacterium group [5'-tggtccgtrtctcagtac] 319-336 The numbers in italics correspond to the nucleotide positions relative to the E. coli 16S rRNA gene. Hybridization buffer: 5x SET,0.2% bovine serum albumin (BSA), 10% dextran sulfate, 0.01% polyadenylic acid, and 0. 1% sodium dodecyl sulfate. Deionized water. Mounting solution (50:50 (v/v) glycerol: I Ox SET). Ecoscint A cocktail (National Diagnostic, Atlanta, GA). Photographic emulsion (type NTB2). Gelatin solution (gelatin solution: 0.02% final concentration gelatin, 0.02% final concentration CrKSO, at a 50:50 (v/v) ratio). Kodak film developer and fixer (Dektol and fixer: Kodak# 146-4114).
Protocol 1. Water samples are collected in 250 ml acid-washed dark bottles. 2. Divide the water sample into four 40 ml equal subsamples in sterile conical tubes. 3. Kill two subsamples by adding 10% formalin and incubate for 1 h at ambient seawater temperature on a rotary shaker. 4. Add tritiated glucose (Dupont) to two replicate samples (final concentration: 10 nM; specific activity: 50 Ci mmole ~;final activity: 10 btCi). 5. Supplement all four of the samples with a tritiated amino acid mixture (final concentration: 5riM; specific activity: 50Cimmole'; final activity: 10 btCi) (Amersham). Incubate at ambient seawater temperature. 6. Sample from all of the live and killed samples over time by withdrawing 2 ml aliquots. 7. Filter the aliquot onto a 25 ram, 0.2 btm Nuclepore polycarbonate filter and rinse the filter four times with 2 ml lx PBS. 8. Measure the radioactivity using a scintillation counter and Ecoscint A cocktail (National Diagnostic, Atlanta, GA). Determine the cell concentration by DAPl staining. When samples reach saturation levels of radioactivity, terminate the uptake of nutrients by adding 10% formalin to live incubations. 10. In a darkroom, melt photographic emulsion (type NTB2) for 1 h at 43°C in a water bath. 11 • Mix the emulsion with gelatin solution at 43°C (gelatin solution: 0.02% final concentration gelatin, 0.02% final concentration CrKSO4 at a 50:50 (v/v) ratio). The emulsion alone has been found to peel off the glass slide during emulsion development o r in situ hybridization. This can be avoided by mixing the emulsion and gelatin solutions together• .
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12. Place an 1TT Night Vision scope and a 15 W safelight approximately 2 m away from all procedures performed in the darkroom. 13. Using the night vision scope to aid seeing in the darkroom, coat the wells on the glass slides containing ten 7-ram-diameter wells by dispensing 20 ~1 of the emulsion:gelatin solution into each well and immediately withdrawing as much solution as possible with a micropipette. Coating the wells separately is necessary to keep the Teflon areas around the wells hydrophobic. Coating the wells individually also prevents the hybridization buffer in one well from merging with buffer from adjacent wells. 14. Allow the slides to dry for 30 rain in total darkness. It is recommended to coat only three slides at one time to minimize background exposure of the emulsion in wells. 15. Set up the filtration tower with 25 ram, 0.2 ~tm Nuclepore filters placed over 0.8 ~tm type AA Millipore filters. 16. Outside the darkroom, under room light, add 10 ml aliquots of the formalin-fixed sample to clean sterile filtration towers. 17. Filter the volume down to approximately 2 ml. 18. Stain the cells with 100 ~1 of DAPI for 10 min in the dark (stock concentration: 0.1 ~tg~ll"). Staining in the dark can be accomplished by covering the filter tower with aluminum foil. 19. Rinse the filters four times with 2 ml lx PBS. Rinsing removes unincorporated radioactive glucose and amino acids. 20. With clean forceps, place the filters, cells side up, onto a drop (10 ~tl) of lx PBS in a Petri dish. 21. Cut the filters into eight equal pieces with a clean and sharp razor blade; carry them to the dark room where they will be transferred to Teflon slides that have been pre-coated. 22. Immediately transfer filters onto coated Teflon slides in the dark room by peeling each of the eight sections of the Nuclepore filter off the Millipore filter and placing it upside down (cells facing down) onto a single well. Only three filters are prepared at any one time to prevent the filters from drying out before being transferred to slides. 23. Allow the slides to dry for 30 min in complete darkness. 24. Place slides in a lightproof box wrapped in aluminum foil and placed in a cardboard box at 4°C for 3 days (for emulsion exposure). 25. Develop the emulsion using Kodak specifications: 2 rain in Dektol developer; 10 s stop in deionized water, 5 min in fixer. 26. Wash the slides with deionized water for 2 rain. 27. Peel off the Nuclepore filters and allow the slides to dry. 28. Hybridize the filters with oligonucleotide probes: (a) Hybridization buffer: 5× SET (lx SET is 150 mM NaC1, 20 mM TrisHC1 pH 7.8, and 1 mM EDTA), 0.2% bovine serum albumin (BSA), 10% dextran sulfate, 0.01% polyadenylic acid, and 0.1% sodium dodecyl sulfate. (b) Probe concentration is 5 ng ml 29. Incubate the slides at 43°C for 3 to 16 h. 30. Rinse the slides with distilled water at 43°C and immerse them three times in 0.2× SET at 43°C for 10 min each time. 190
31. Allow the slides to air dry. 32. Mount the slides with glycerol: 10x SET (50:50 (v/v)) and store at -20°C for at least 1 h. 33. Detect probe hybridization with fluorescence and uptake of radiolabeled substrates by transmitted light microscopy. (a) Total DAPI counts with UV excitation (fluorescence microscope equipped with a UV filter (Olympus type U-MWU)). (b) Probe fluorescence counts with green excitation (fluorescence microscope equipped with a chroma type TRITC U-M41002 filter). (c) Microautoradiography counts represented by small dark silver grains (transmitted light microscopy). (d) Counts for cells labeled with fluorescent probes and simultaneously labeled with microautoradiography. An example of microautoradiographs generated using the MICRO-FISH method (Cottrell and Kirchman, 2000) is shown in Plate 3. The MICROFISH method identifies bacterial cells that incorporate tritiated compounds by the presence of silver grains adjacent to the blue DAPIstained cells (Panel A). The phylogenetic classification of bacterial cells is determined by hybridization with 16S rRNA oligonucelotide probes conjugated to Cy 3 (Panel B).
Advantages Simultaneously determines the phylogenetic identity and specific metabolic activity of individual cells. Results from the triple labeling STARFISH protocol are consistent with standard fluorescent in situ hybridization (FISH) protocols. The scope of the technique application is broad with respect to the identity of target microorganisms and metabolic activity of interest.
o
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°4 Disadvantages
e" D.
•
Presence of silver grains in the photographic emulsion that do not correspond to a cell in either the DAPI or probe fields may result in ambiguous analysis. • Slow uptake and incorporation of labeled substrates may result in cells appearing active in the probe field (based on rRNA content) and not active in the autoradiographic emulsion. • Procedure is tedious and labor intensive.
4,41,4,~,4,~ L I V E / D E A D B A C L I G H T VIABILITY KIT
TM
BACTERIAL
Introduction Microbial viability can be assessed by monitoring biological factors that are altered during loss of viability including variations in membrane permeability (Jepras et al., 1995). A bacterium is assumed to be viable and 191
potentially active if the membrane is not damaged and assumed to be dead if the membrane is compromised (Decamp and Rajendran, 1998a,b). Membrane integrity analysis is based on the capacity of the cells to exclude compounds, such as fluorescent intercalating dyes, which when used at low concentrations do not normally cross intact membranes (Jepras et al., 1995). A membrane-impermeant stain that can passively diffuse through a cell wall can act as an indicator of loss in membrane integrity and thus as an indicator of cell viability. The LIVE/DEAD BacLight'" bacterial viability kit (Molecular Probes, Oregon) provides a two-color fluorescent assay of bacterial viability. The kit utilizes mixtures of two nucleic acid stains: SYTO 9 and Propidium Iodide (PI). These stains differ in their spectral characteristics and in their ability to penetrate healthy bacterial cells. SYTO 9 is a green fluorescent stain that generally stains all bacteria in a population regardless of their viability (Molecular Probes, Product Information). Propidium iodide is a membrane-impermeant stain that penetrates only bacteria with damaged membranes (Jernaes and Steen, 1994; Lopez-Amoros et al., 1995; Comas and Vives-Rego, 1997; Williams et al., 1998). When combined, the two stains rapidly distinguish between live bacteria with intact membranes from dead bacteria with compromised membranes. Although not initially used with marine samples for technical reasons, recently the BacLight" protocol has been modified and applied with apparent success to marine samples (Decamp and Rajendran, 1998a,b; Gasol et al., 1999). In addition, because of its simplicity and potential applicability with flow cytometry and epifluoresence microscopy (Molecular Probes, Product Information), the LIVE/DEAD BacLighV" kit remains an attractive methodology for exploring the physiological status of marine bacteria.
Principle SYTO 9 is a nucleic acid stain that penetrates the cell wall of intact live cells. The dye can stain both DNA and RNA and is permeable to virtually all cell membranes. SYTO 9 accumulates inside all cells and can be visualized with Narrow Blue (NB) excitation (Molecular Probes, Product Information). Propidium iodide is a fluorescent nucleic acid dye, which stains by intercalating into nucleic acid molecules (Jepras et al., 1995; Lopez-Amoros et al., 1995, 1997). PI binds to both DNA and RNA and is non-specific with respect to base sequence. PI accumulates inside cells that have compromised or leaky membranes and stains the nucleus light to dark red in color when visualized with Wide Green (WG) excitation (Williams et al., 1998). Accumulation of PI inside the cell causes a reduction in the SYTO 9 stain fluorescence when both dyes are present. Propidium iodide competes for nucleic acid binding sites with SYTO 9 and displaces the already bound dye (Virta et al., 1998). With an appropriate mixture of SYTO 9 and PI stains, bacteria with intact membranes stain green, whereas bacteria with damaged membranes stain fluorescent red (DeCamp and Rajendran, 1998a,b). These two stains when used in combination can be visualized simultaneously or separately.
192
Equipment and reagents • Vacuum manifold, 25 mm vacuum filter holders. • Epifluorescence microscope equipped with a Narrow Blue (NB) and aWide Green (WG) filter set. • Vacuum source. • 1.5 ml Eppendorf tubes. • 0.2 Mm black polycarbonate filters (Micron Separations, Inc). • Distilled water. • Formalin. • LIVE/DEAD BacLight ~' Kit: (a) ComponentA: 300 MI solution (in DMSO) of 3.34 mM SYTO 9 dye. (b) Component B: 300 MI solution (in DMSO) of 20 mM Propidium Iodide. (c) Component C: BacLight mounting oil; I 0 ml, for bacteria immobilized on membranes. Refractive index at 25°C is 1.517+0.0003. NOT IMMERSION OIL. The manufacturer recommends the following: • These components should be stored frozen a t - 2 0 ° C and protected from light. • The reagents should be warmed to room temperature and centrifuged briefly each time they are removed from the freezer. • Vials should always remain tightly sealed. Under these conditions the components are stable for at least one year. • Glass microscope slides. • Glass 18 mm square coverslips.
o
Protocol 1. Water samples collected in the field can be fixed and stored in 4% final concentration of buffered formalin and stored at 4°C in the dark until analyzed. 2. Bacterial suspensions: remove traces of growth media before staining bacteria with kit reagents. Nucleic acids and other media components can bind the dyes in unpredictable ways. A single wash step is usually sufficient to remove significant traces of interfering media components. Phosphate buffers are not recommended because they appear to decrease staining efficiency (Haugland, 1996). 3. Combine equal volumes of Component A and Component B in a microfuge tube and mix thoroughly. 4. Add 3 ~tl of the dye mixture for each 1 ml of the bacterial suspension or aquatic sample. When used at the recommended dilutions, the reagent mixture will contribute 0.3% DMSO to the staining solution. Higher DM50 concentrations may adversely affect staining. 5. Mix thoroughly and incubate at room temperature in the dark for 15 rain. 6. Filter the suspension through a 0.2~tm black polycarbonate filter placed onto a manifold set-up. Wash with 2ml of distilled water, mount onto a glass microscope slide with the mounting oil supplied with the BacLight kit, OR
193
i~00) m
efl.
U
Trap 5 gl of the stained bacterial suspension between a slide and an 18 m m square coverslip. . E n u m e r a t e stained cells with an epifluorescence microscope e q u i p p e d with a N a r r o w Blue (NB) filter set to visualize SYTO-9-stained cells and a Wide Green (WG) filter set to visualize PI-stained cells. Molecular probes r e c o m m e n d s the use of O m e g a Optical filter sets (for SYTO 9 - - O-5715 or O-5716; for PI - - 0-5723, 0-5724, 0-5733) which are available through their catalog.
Advantages • All cells are visualized, not restricted to specific groups of bacteria. • SYTO 9 dye coupled with PI allows for cells to be visualized as dead or live simultaneously. • Both SYTO 9 and PI express fluorescence enhancement upon binding nucleic acids making the differentiation of live and dead cells uncomplicated. • The BacLight T~'kit can be coupled with flow cytometry. • Rapid and simple protocol.
Disadvantages • Technique provides information about potential viability only, not activity of cells. • BacLight Tr' is unable to indicate dead cells that have already lost their cytoplasmic content (Lawrence et al., 1997). • Relies on a single cellular criterion. • Staining by PI and SYTO 9 is highly dependent on salt concentration. Haugland (1996) reported that BacLight '~ may not be suitable for marine samples although it has been used successfully in some marine environments (Decamp and Rajendran, 1998a, b; Gasol et al., 1999).
eeeeee
DIRECT VIABLE COUNTING METHOD
[DVC]: K O G U R E
Introduction A cell that is actively g r o w i n g and dividing can be considered viable and active. Thus, the ability of a cell to g r o w and divide can be used as a p r i m a r y characteristic of living, active cells (Joux and LeBaron, 1997). During the late 1970s, Kogure et al. (1979) d e v e l o p e d the Direct Viable Counting (DVC) m e t h o d to identify and e n u m e r a t e cells that were actively growing. This m e t h o d utilizes the antibiotic nalidixic acid to interfere with normal cell division. In the presence of this antibiotic, sensitive cells g r o w but do not divide and can therefore be easily identified microscopically as elongated or enlarged cells.
194
One of the limitations in applying the original Kogure DVC method (1979) to complex communities is the presence of bacteria in the environment that are resistant to nalidixic acid and are therefore able to grow and divide normally in the presence of this antibiotic. This limitation has led to modifications of the initial procedure (Kogure et al., 1984, 1987; Tabor and Neihof, 1984; Coallier et al., 1994). Recent improvements have utilized multiple antibiotic cocktails that act similarly to nalidixic acid (Kogure ct tTl., 1984; Servis et al., 1993; Joux and LeBaron, 1997). In general DVC results are well correlated with other measures of cell activity, i.e. cell reducing potential (Maki and Remsen, 1981; Tabor and Neihof, 1984).
Principle Nalidixic acid (nal) is an antibiotic that affects the cell by inhibiting the bacterial enzymes DNA gyrase and topoisomerase II by binding to the DNA gyrase complex (Prescott et a[., 1997). DNA gyrase inhibition disrupts DNA synthesis and replication due to improper coiling of the DNA molecule. While DNA synthesis is disrupted by nal, synthesis of RNA, protein, and cellular components continue (Maki and Remsen, 1981). Active cells that are unable to divide will continue to grow and become elongated. The visible enlargement of the cell is the basis for recognizing active bacteria. Other antibiotics that have been used with this method include piromidic acid, pipemidic acid, ciproflaxin, and cephalexin (Joux and LeBaron, 1997). The first three antibiotics listed inhibit DNA gyrase as well, however, cephalexin inhibits the transpeptidase and transglycosylase reactions, which serve to cross-link the cell wall in bacteria. Cephalexin blocks the formation of the septum during cell division and in st) doing, produces elongated cells. These antibiotics working together have been found to inhibit 96(7~: of all isolates tested (Joux and LeBaron, 1997).
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Equipment and reagents
•
•
Epifluorescent microscope equipped with a UV filter set. Vacuum manifold, 2_5 mm vacuum filter holders. Vacuum source. 20°C incubator. Sterile yeast extract solution (final concentration: 50 mg ml '). 37% pro-grade formalin. Antibiotics: see Table 10.2_. DAPI (stock concentration: 2_00 ~tg ml '). Sigma Chemical (St Louis, MO). DAPI stock solutions are dissolved in distilled water and can be stored in the refrigerator at 2 ~ ° C in the dark until needed. Acridine Orange (final concentration: 0.01%). Sigma (St Louis, M O ) . A O stock solutions are dissolved in distilled water and can be scored in the refrigerator at 2-4°C in the dark until needed. 0.5 M NaOH.
195
Table 10.2 List of the antibiotics used for inhibiting cell division with the direct viable counting (DVC) technique.These antibiotic solutions are filter sterilized through 0.2 ~tm membrane filters and can be pre-made and stored in the dark at 4°C Antibiotic
Stock conc.
Final conc.
Solvent
Nalidixic acid
500 big ml '
20 ~g ml ~
Pipemidic acid
500 ~lg ml '
10 btg ml '
Piromidic acid
500 btg ml '
10 btg mt '
Cephalexin
500 btg ml '
10 btg ml '
Ciproflaxin
500 ~lg ml ~
0.5 btg ml '
Sterile water Sterile water Sterile water Sterile water Sterile water
• • • • • • • •
Distributor
MilliQ MilliQ MilliQ MilliQ MilliQ
Sigma St Louis, MO Sigma St Louis, MO Sigma St Louis, MO Sigma St Louis, MO Bayer Pittsburgh, PA
0.2 IJm filtered distilled water. 25 mm, 0.2/Jm Nuclepore black polycarbonate filters. Screw cap glass bottles (amber or clear). 15 ml glass tubes (test tubes) and caps. Glass microscope slides. Glass microscope cover slips. Microscope grade immersion oil. Non-fluorescing immersion oil.
Protocol
1. Samples are collected and placed into screw cap amber or clear glass bottles. 2. Fill sterile 15 ml glass tubes with 9 ml of sample (duplicate samples are recommended). 3. For each sample, a control (1-2 ml d e p e n d i n g on the bacterial biomass of the sample) is fixed in f o r m a l d e h y d e (2% final concentration with 37% formalin) and stained as described below in step 8. 4. Aseptically enrich each of the samples with 1 ml of yeast extract (final concentration: 50 mg 1 ~) 5. Add the antibiotic cocktail; see Table 10.2. 6. Incubate the samples at 20 ° C for 6 to 8 h. 7. Fix the samples with 0.2~7, f o r m a l d e h y d e (final concentration). Fixing the samples terminates synthesis of cellular materials and growth. 8. Stain cells with Acridine Orange (AO). Stain cells with a 0.01% final concentration of AO. Mix 1 ml of the water sample with 1 ml of dilution water and 0.2 ml of 0.1% AO (stock solution). Vortex and incubate for 2 minutes, OR 4'6-diamidino-2-phenylindole (DAPI). Stain cells with 2.5 ~g of DAPI per 1 ml of sample. Vortex and incubate the sample in the dark for 30 rain. 196
9. Place a 25 mm, 0.2 pm Nuclepore black polycarbonate filter onto a tower set-up. Pipette solution of cells and stain onto a filter tower and wash the glass bottle with 2 ml of filtered water (0.2 ~tm pore-size filter). Pipette this volume onto the filter tower and filter until dry with gentle vacuum. 10. Remove the filter from the column and place it onto a glass slide, cell side up. 11. Place a drop of non-fluorescing immersion oil on top of the filter and cover with a coverslip. 12. Enumerate the total cells (AO or DAPI) and active cells (elongated) using an epifluorescent microscope equipped with a blue excitation filter and UV filter set, respectively. Joux and LeBaron (1997) recomm e n d the use of a Zeiss UV filter set (4877-02). Excitation 365 nm, barrier 420 nm. An example of bacteria cells before and after incubation with a combination of antibiotics (nal, pipemidic acid, piromidic acid, cephalexin, and ciproflaxin) that inhibit cell division is presented in Figure 10.2.
Advantages The assessment of viability is based on indicators of DNA, RNA, and protein synthesis and not on the concentration of those molecules within the cell. Thus, cells must be actively growing and dividing in order to be detected. The DVC method is theoretically not restricted to particular cell types. Protocol is easy to perform, and has a short incubation time. i
Disadvantages •
• •
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Technique necessitates the in vitro growth of cells and therefore is subject to cultivation bias. The extent of this bias is unknown and likely to be variable between samples. The addition of a growth substrate (e.g. yeast extract) may introduce artefacts associated with cultivation. The natural occurrence of antibiotic resistant cells in natural samples may bias results.The extent of this bias is unknown and likely varies among samples.
CONCLUSIONS In this chapter we have described five methods available to assess the physiological status of individual cells in marine water samples. A sixth method 'CTC' is described elsewhere in this volume (Sherr et al., Chapter 8). The general advantages and disadvantages of each of these methods are summarized in Table 10.3. In general, we have considered and compared methods based on their simplicity, reliability, specificity, versatility, and the degree that they have been used in published reports. 197
°4 ca.
198
Table 10.3 Summary of technical aspects of the five methods employed for determining the physiological status of bacteria, described in detail in the text
VSP
Microautoradiography Nudeoid and probes
Kogure
Live/Dead
Simplicity
Intermediate 2 hours
Difficult ~ 2 days
Simple < 1 hour
Simple ~ 10 hours
Simple < 1 hour
Versatility, reliability
Good Two independent criteria to assess the physiological status
Good Two independent criteria to assess the physiological status
Fair One criteria
Fair One criteria
Fair One criteria
Cell specificity
Total community or specific groups of bacteria
Total community or specific groups of bacteria
Total community
Total Total community community
Key references
° This chapter * Williams et al., 1998
• Ouverney and Fuhrman, 1999 • Cottrell and Kirchman, 1999
• Choietal., * J o u x a n d *Gasolet 1996 LeBaron, al., 1999 • Zweifel 1997 ° DeCamp and ° Kogure and Hagstrom, et al., 1987 Rajendran, 1995 1998 a,b
The d e v e l o p m e n t of n e w m e t h o d o l o g i e s in the field of microbial ecology has b e c o m e a science unto itself (Paul, 1993). A s p e c t r u m of alternative methodological a p p r o a c h e s is n o w either u n d e r d e v e l o p m e n t or available for m e a s u r i n g single cell attributes of aquatic bacterioplankton. These techniques and protocols target specific cellular criteria that can be associated with the physiological status of an individual bacterial cell. As these m e t h o d s are d e v e l o p e d , used, and c o m p a r e d in various natural systems, it is clear that different m e t h o d s can yield different estimates of the p r o p o r t i o n of bacterial cells that are alive, viable and metabolically active (Choi et al., 1999). The active fraction of bacterial p o p u l a t i o n s that h a v e been reported in the literature to date range from nearly 0% to nearly 100% d e p e n d i n g on the e n v i r o n m e n t and the technique used to estimate this parameter. H o w e v e r , e m e r g i n g from these disparate data sets is the general conclusion that, in m o s t marine e n v i r o n m e n t s and at any given point in time, a significant fraction of the bacteria e n u m e r a t e d b y direct microscopic counting m e t h o d s are either dead or inactive. To complicate these conclusions there is a relative sparseness of studies that directly c o m p a r e m e t h o d s in a systematic fashion. Karner and F u h r m a n (1997) conducted a rare e x a m p l e of such a study. These investigators c o m p a r e d
F i g u r e 10.2. Fluorescence micrographs of DAPI-stained bacteria from a marine sample before incubation (A), after 6 h incubation with nalidixic acid (20 gg ml ~) (B), and after 18 h incubation with the antibiotic cocktail (C). Substrate responses are identified by arrows. Bar, 10 btm.
199
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results obtained from a variety of m a r i n e w a t e r s a m p l e s using the CTC, nucleoid staining, microautoradiography, 16S r R N A oligonucleotide probes, and DAPI staining methods. Although the results obtained from these c o m p a r i s o n s varied, based on their s t u d y Karner and F u h r m a n reached the conclusion that m i c r o a u t o r a d i o g r a p h y and ribosomal RNA targeted probes were m o s t likely the m o s t reliable indicators of physiological activity since these two m e t h o d s were self-consistent. Extending this logic, it seems reasonable that the m o s t useful methodological a p p r o a c h for estimating the physiological status of an individual cell will combine c o m p o n e n t s that s i m u l t a n e o u s l y target multiple cellular targets that can act as indicators of physiological status. Of the m e t h o d s described in this chapter, several allow the simultaneous evaluation of multiple criteria of physiological status (Williams et al., 1998; Lee et al., 1999; O u v e r n e y and F u h r m a n , 1999; Cottrell and Kirchman, 2000). Therefore, these m e t h o d s are considered to be the m o s t reliable m e t h o d s available today. However, all of the m e t h o d s available h a v e limitations and therefore results m u s t be interpreted cautiously and with respect to the specific cellular criteria examined. Future m e t h o d d e v e l o p m e n t should concern itself with m e t h o d s that are straightforward, simple to p e r f o r m and reproduce, and h a v e universal application to a variety of cell types. The need for these d e v e l o p m e n t s is undeniable.
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Comas, J. and Vives-Rego, J. (1997). Assessment of the effects of gramicidin, formaldehyde, and surfactants on Escherichia coli by flow cytometry using nucleic acid and membrane potential dyes. Cytometry 29, 58-64. Cottrell, M. T. and Kirchman, D. L. (2000). Natural assemblages of marine proteobacteria and cytophaga-flavobacter consuming low and high-molecular weight dissolved organic matter. Appl. Envilvn. Microbiol. 66, 1692-1697. Decamp, O. and Rajendran, N. (1998a). Assessment of bacterioplankton viability by membrane integrity. Mar. Pollut. Bull. 36, 739-741. Decamp, O. and Rajendran, N. (1998b). Bacterial loss and degradation of bacterial membrane in preserved seawater samples. Mar. Pollut. Bull. 36, 856-859. Del Giorgio, P. A. and Scarborough, G. (1995). Increase in the proportion of metabolically active bacteria along gradients of enrichment in freshwater and marine plankton: implications for estimates of bacterial growth and production. J. Phmkton Res. 17, 1905-1924. DeLong, E. E (1993). Single-cell identification using fluorescently labeled ribosomal RNA-specific probes. In: Handbo~)k of Methods in Aquatic Microbial Ecology (Kemp, P. E, Sherr, B. E, Sherr, E. B. and Cole, J. J. Eds), pp. 285-294. Lewis Publishers, London. DeLong, E. E, Wickham, G. S. and Pace, N. R. (1989). Phylogenetic stains: ribosomal RNA-based probes for identification of single cells. Science 243, 1360-1363. Ducklow; H. W. (1999). The bacterial component of the oceanic euphotic zone. FEMS Microbiol. Ecol. 30, 1-10. Epstein, S. S. and Rossel, J. (1995). Methodology of iu situ grazing experiment: evaluation of a new vital dye for preparation of fluorescently labeled bacteria. Mar. Ecol. ProS. Ser. 128, 143-150. Frischer, M. E., Floriani, P. J. and Nierzwicki-Bauer, S. A. (1996). Differential sensitivity of 16S rRNA targeted oligonucleotide probes for fluorescence in situ hybridization is a result of ribosomal higher order structure. Call. J. Microbiol. 42, 1061 1071. Gasol, J. M., Zweifel, U. L., Peters, E, Fuhrman, J. A. and Hagstrom, A. (1999). Significance of size and nucleic acid content heterogeneity as measured by flow cytometry in natural planktonic bacteria. Appl. EHviro11. Microbiol. 65, 4475-4483. Giovannoni, S. J., DeLong, E. E, Olson, G. J. and Pace, N. R. (1988). Phylogenetic group-specific oligodeoxynucleotide probes for identification of single microbiaI cells. J. Bactcriol. 170, 720-726. Glockner, E O., Amann, R. I., Alfreider, A., Pernthaler, J., Psenner, R., Trebesius, K. and Schleifer, K.-H. (1996). An in situ hybridization protocol for detection and identification of planktonic bacteria. Syst. Appl. Microbiol. 19, 403-406. Haugland, R. P. (1996). Moh'culnr Pr(~bes Haluibook off FhloresceiH Probes and Research Chemicals. Molecular Probes, Eugene, OR. Heissenberger, A., Leppard, G. G. and Hemdl, G. J. (1996). Relationship between the intracellular integrity and the morphology of the capsular envelope in attached and free-living marine bacteria. Appl. Enviro11. Microbiol. 62, 4521-4528. Hicks, R. E., Amann, R. I. and Stahl, D. A. (1992). Dual staining of natural bacterioplankton with 4',6-diamidino-2-phenylindole and fluorescent oligonucleotide probes targeting kingdom-level 16S rRNA sequences. Appl. EnviroH. Microbiol. 58, 2158-2163. Hobble, J. E., Daley, R. J. and Jasper, S. (1977). Use of Nuclepore filters for counting bacteria by epifluorescence microscopy. Appl. E~viron. Microbiol. 33, 1225-1228. Hong, Y, Frischer, M. E., Verity, P. G. and Danforth, J. D. (1998). In vivo rRNA degradation rates: identification of dead but recently active cells in marine microbial communities. American Society for Microbiology General Meeting. Atlanta, GA, N-120.
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°2 e-
Hoppe, H. G. (1976). Determination and properties of actively metabolizing heterotrophic bacteria in the sea investigated by means of micro-autoradiography. Mar. Biol. (Berlin) 36, 291-302. Jepras, R. I., Carter, J., Pearson, S. C., Paul, E E. and Wilkinson, M. J. (1995). Development of a robust flow cytometric assay for determining numbers of viable bacteria. Appl. Environ. Microbiol. 61, 2696-2701. Jernaes, M. W. and Steen, H. B. (1994). Staining of Escherichia coli for flow cytometry: influx and efflux of ethidium bromide. Cytometry 17, 302-309. Joux, E and LeBaron, R (1997). Ecological implications of an improved direct viable count method for aquatic bacteria. Appl. Environ. Microbiol. 63, 3643-3647. Karner, M. and Fuhrman, J. A. (1997). Determination of active marine bacterioplankton: a comparison of universal 16S rRNA probes, autoradiography, and nucleoid staining. Appl. Enviroil. Microbiol. 63, 1208-1213. Kepner, R. L., Jr. and Pratt, J. R. (1994). Use of fluorochromes for direct enumeration of total bacteria in environmental samples: past and present. Microbiol. Rev. 58, 603-615. Kogure, K., Simidu, U. and Taga, N. (1979). A tentative direct microscopic method for counting living marine bacteria. Can. 1. Microbiol. 25, 415-420. Kogure, K., Simidu, U. and Taga, N. (1984). An improved direct viable count method for aquatic bacteria. Arch. Hydrobiol. 102, 117-122. Kogure, K., Simidu, U., Taga, N. and Colwell, R. R. (1987). Correlation of direct viable counts with heterotrophic activity for marine bacteria. Appl. Environ. Microbiol. 53, 2332-2337. Lane, D. J., Pace, B., Olsen, G. J., Stahl, D. A., Sogin, M. L. and Pace, N. R. (1985). Rapid determination of 16S ribosomal RNA sequences for phylogenetic analyses. Proc. Natl. Acad. Sci. USA 82, 6955-6959. Lawrence, J. R., Korber, D. R., Wolfaardt, G. M. and Caldwell, D. E. (1997). Analytical imaging and microscopy techniques. In: Manual of Enviromnental Microbiology (C. J. Hurst, G. R. Knudsen, M. J. McIerney, L. D. Stetzenbach and M. V. Walter, Eds), pp. 29-51. American Society for Microbiology Press, Washington, DC. Lee, N., Nielsen, R H., Andreasen, K. H., Juretschko, S., Nielsen, J. L., Schleifer, K. H. and Wagner, M. (1999). Combination of fluorescent iH situ hybridization and microautoradiography - - a new tool for structure-function analyses in microbial ecology. Appl. Environ. Microbiol. 65, 1289-1297. Lee, S. and Kemp, R E (1994). Single-cell RNA content of natural marine planktonic bacteria measured by hybridization with multiple 16S rRNA-targeted fluorescent probes. Limnol. Oceam~gr. 34, 869 879. Lee, S., Malone, C. and Kemp, R E (1993). Use of multiple 16S rRNA targeted fluorescent probes to increase signal strength and measure cellular RNA from natural planktonic bacteria. Mar. Ecol. Prog. Ser. 101, 193-201. Lopez-Amoros, R., Comas, J. and Vives-Rego, J. (1995). Flow cytometric assessment of Escherichia coil and Salmonella typhimurium starvation-survival in seawater using rhodamine 123, propidium iodide, and oxonol. Appl. Environ. Micn)biol. 61, 2521-2526. Lopez-Amoros, R., Castel, S., Coams-Riu, J. and Vives-Rego, J. (1997). Assessment of E. coli and Sahnonella viability and starvation by confocal laser microscopy and flow cytometry using rhodamine 123, DiBAC4(3), propidium iodide, and CTC. Cytometry 29, 298-305. Lovejoy, C., Legendre, L., Klein, B., Tremblay, J.-E., Ingrain, R. G. and Therriault, J.-C. (1996). Bacterial activity during early winter mixing (Gulf of St Lawrence, Canada). Aquatic Microbial Ecol. 10, 1-13.
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Lundgren, B. (1981). Fluorescein diacetate as a stain of metabolically active bacteria in soil. Oikos 36, 17-22. Maki, J. S. and Remsen, C. C. (1981). Comparison of two direct-count methods for determining metabolizing bacteria in freshwater. Appl. Environ. Microbio[. 41, 1132-1138. Mason, D. J., Lopez-Amoros, R., Allman, R., Stark, J. M. and Lloyd, D. (1995). The ability of membrane potential dyes and calcafluor white to distinguish between viable and non-viable bacteria. J. Appl. Bacteriol. 78, 309-315. Meyer-Reil, L. A. (1978). Autoradiography and epifluorescence microscopy combined for the determination of number and spectrum of actively metabolizing bacteria in natural waters. Appl. Environ. Microbiol. 36, 506-512. Molecular Probes, Inc. (1997). Product Information Sheet. LIVE/DEAD BacLigh.~ Bacterial Viability Kits. Molecular Probes, inc., Eugene, Oregon. Ouverney, C. C. and Fuhrman, J. A. (1999). Cornbined microautoradiography-16S rRNA probe technique for determination of radioisotope uptake by specific microbial cell types in situ. Appl. E1~viro~l. Microbiol. 65, 1746-1752. Paul, J. H. (1993) The advances and limitations of methodology. In: Aquatic Microbiology alz Eco[o~¢ical Approach (T. E. Ford, Ed.,), pp. 15-46. BlackweI1 Scientific, Boston. Prescott, L. M., Harley, J. P. and Klein, D. A. (1997). Microbiology, 2nd edn. Win. C. Brown Publishers, Dubuque, Iowa. Proctor, L. M. and Fuhrman, J. A. (1990). Viral mortality of marine bacteria and cyanobacteria. Nature 343, 60 62. Rodriguez, G. G., Phipps, D., lshiguro, K. and Ridgway, H. E (1992). Use of a fluorescent redox probe for direct visualization of actively respiring bacteria. Appl. Enviro11. Microbiol. 58, 1801-1808. Schaechter, E., Maaloe, O. and Kjeldgaard, N. O. (1958). Dependence on medium and temperature of cell size and chemical composition during balanced growth of Salmonella typhimurium. J. GeH. Microbiol. 19, 592-606. Servis, N. A., Lytle, M. S., Midthun, D. B., Leake, R. A. and Adams, J. C. (1993). Comparison of isopropyl cinodine with nalidixic acid in the direct viable count. J. Appl. Bocteriol. 75, 583-587. Sherr, B. E, del Giorgio, P. and Sherr, E. B. (1999). Estimating abundance and single cell characteristics of actively respiring bacteria via the redox dye, CTC. Aquatic Microbiol Ecol. 18, 117-131. Sherr, E. B., Sherr, B. E and Sigmon, C. T. (2000). Activity of marine bacteria under incubated and in situ conditions. Aquatic Micn~biol. Ecol. 20, 213-223. Shopox, A., Williams, S. C. and Verit}; P. G. (2000). Image analysis for the discrimination and enumeration of bacteria and viruses in aquatic samples. Aquatic Microbiol Ecol. 22, 103-111. Sieracki, M, E., Reichenbach, S. E. and Webb, K. L. (1989). Evaluation of automated threshold selection methods for accurately sizing microscopic fluorescent cells by image analysis. Appl. EnviroH. Microbiol. 55, 2762. Spring, S., Amann, R., Ludwig, W., Schleifer, K. H., Gemerden, H. and Peterson, K. (1993). Dominating role of an unusual magnetotactic bacterium in the micro aerobic zone of a freshwater sediment. Appl. Environ. Microbiol. 59, 2397 2403. Tabor, P. S. and Neihot, R. A. U984). Direct determination of activities for microorganisms of Chesapeake Bay populations. Appl. Environ. Microbiol. 48, 1012-1019. Tsuji, T., Kawasaki, Y., Takeshima, S., Sekiya, T. and Tanaka, S. (1995). A new fluorescence staining assay for visualizing living microorganisms in soil. Appl. El~viroll. Microbiol. 61, 3415-3421.
203
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Vescio, P. A. and Nierzwicki-Bauer, S. A. (1995). Extraction and purification of PCR amplifiable DNA from lacustrine subsurface sediments. J. Microbiol. Methods 21, 225-233. Virta, M., Lineri, S., Kankaanpaa, P., Karp, M., Peltonen, K., Nuutila, J. and Liluis, E. M. (1998). Determination of complement-mediated killing of bacteria by viability staining and bioluminescence. Appl. Environ. Microbiol. 64, 515-519. Vosjan, J. H. and van Noort, G. J. (1998). Enumerating nucleoid-visible marine bacterioplankton: bacterial abundance determined after storage of formalin fixed samples agrees with isopropanol rinsing method. Aquatic Microbial Ecol. 14, 149-154. Whitman, W. B., Coleman, D. C. and Wiebe, W. J. (1998). Prokaryotes: the unseen majority. Prec. Natl. Acad. Sci. USA 95, 6578-6583. Williams, S. C., Hong, Y., Danavall, D. C. A., Howard-Jones, M. H., Gibson, D., Frischer, M. E. and Verity, P. G. (1998). Distinguishing between living and nonliving bacteria: evaluation of the vital stain propidium iodide and its combined use with molecular probes in aquatic samples. I. Microbiol. Methods 32, 225-236. Wolfe, S. L. (1983). Introduction to Cell Biology. Wadsworth Publishing Co., Belmont, CA. Zimmermann, R., Iturriaga, R. and Becker-Birck, J. (1978). Simultaneous determination of the total number of aquatic bacteria and the number thereof involved in respiration. Appl. Environ. Microbiol. 36, 926-935. Zweifel, U. L. and Hagstrom, A. (1995). Total counts of marine bacteria include a large fraction of non-nucleoid containing bacteria (ghosts). Appl. Environ. Mictvbiol. 61, 2180-2185.
List of suppliers Amersham Pharmacia Biotech 800 Centennial Avenue, PO Box 1327, Piscataway, NJ 08855, USA; 1-800-526-3593
Fuller D'Albert, Inc. PO Box 2706, Fairfax, VA 22031, USA; 1-703-591-8000
Radioactive supplies
Radioactive supplies
Bayer 100 Bayer Rd, Pittsburgh, PA 15205, USA; 1-412-777-2000
Gelman Sciences Corp (Pall Gelman) 600 South Wagner, Road Building 3, Ann Arbor, M148103-9019, USA; 1-800-521-1520
Antibiotics
M e m b r a n e filters
Dupont/New England Nuclear 549 Albany Street, Boston, MA 02118, USA; 1-800-551-2121 Radioactive supplies
Integrated DNA Technologies 1710 Commercial Park, Coralville, IA 52241, USA; 1-800-328-2661 Oligonucleotides
204
Millipore 8(9 Ashby Road, Bedford, MA 01730, USA; 1-8(]0-645-5476
Photometrics Ltd 3440 E Britannia Drive, Tucson, AZ 85706, USA; 1-520-889-9933
Membrane filters
Digital cameras and image analysis equipment
Molecular Probes Inc. 4849 Pitchford Avenue, Eugene, OR, 97402, USA; 1-541-465-8300
LIVE/DEAD BacLight Kit National Diagnostic 305 Patton Drive, Atlanta, GA 30336, USA; 1-800-526-3867
Radioactive supplies Nuclepore Corporation 7035 Commerce Circle, Pleasanton, CA 94566, USA
Membrane filters
Photonic Science Limited Millham, Mountfield, Robertsbrid~,,e, East Sussex, TN32 5LA UK; +44 (0) I580 881 199
Digital cameras and image analysis equipment Research Organics 4353 E 49th Street, Cleveland, OH 44125, USA; 1-800-321-0570
FITC, FluorX, TRITC Sigma Chemical Company PO Box 14508, St. Louis, MO 63178, USA; 1-800-325-3010 O
Olympus America Incorporated 4 Nevada Drive, Lake Success, N Y 11042, USA
Epifluorescence microscopes Osmonics/Micron Separations Incorporated 135 Flanders Road, PO Box 1046, Westboro, MA 01581, USA; 1-800-444-8212
Membrane filters
DAPI, Acridine Orange, Propidium Iodide, Antibiotics
m
Southern Micro Instruments Atlanta, GA 30339, LISA; 1-800-241-3312
.~ "F.
Epifluorescence microscopes
ca.
Whatman 9 Bridewell Place, Clifton, NJ 07014, USA; 1-800-922-0361
Membrane filters Other supplies including glassware, slides, slide boxes, coverslips, pipette tips, eppendorf tubes, conical tubes, and coplin jars can be obtained from any large scientific supply company including Fisher Scientific and VWR Scientific. The research-grade chemicals (formaldehyde, ethanol, Tris, EDTA, MgSO,, 2-propanol, BSA, Polyadenylic acid, dextran sulfate) can be obtained from Sigma Aldrich Chemical Company.
205
i/1
ACKNOWLEDGMENTS This work was supported by grants from the National Science foundation to M.E.E and P.G.V. OCE-9617884 and OCE-9906734, and a National Science Foundation CIRE Activity award OCE-9872694 also to P.G.V. and M.E.E The Department of Energy also provided support for this work DEFG02-88ER62531. We wish to acknowledge the important contribution of Ms. Samanthia Williams for the initial development of the VSP method and Y. Hong, J. Danforth, A. Shopov, and V. Ballard for technical assistance. We especially thank L. Cowden for critically critiquing the manuscript and A. Boyette for preparing the figures.
206
11 Fluorescence in situ Hybridization (FISH) with rRNA-targeted Oligonucleotide Probes Jakob Pernthaler', Frank-Oliver GI6ckner',Wilhelm Sch6nhuber 2 and RudolfAmann' 'Max-Planck-lnstitute for Marine Microbiology, Celsiusstrasse I, 28359 Bremen, Germany; 2Institute Pasteur, Physiologie Microbienne, 28, Rue du Docteur Roux, 75724 Paris CEDE)( 15, France
CONTENTS Introduction FISH of environmental samples on membrane filters
Probe design and testing Applications Conclusions ,,~ ¢.
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INTRODUCTION
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Fluorescence in situ hybridization (FISH) with rRNA-targeted probes is, among other things, a staining technique that allows phylogenetic identification of bacteria in mixed assemblages without prior cultivation (Plate 4) by means of epifluorescence and confocal laser scanning microscopy, or by flow cytometry (Giovannoni et al., 1988; DeLong et al., 1989; Amann et aI., 1990a; 1990b; 1996). FISH with polynucleotide DNA probes, and FISH with oligonucleotide probes targeted to mRNA has also been described (Trebesius et al., 1994; Wagner et al., 1998; DeLong et al., 1999). This protocol will, however, exclusively focus on FISH with oligonucleotide probes for the purpose of bacterial identification, i.e. to analyze bacterial community structure, and to follow the spatial and temporal dynamics of individual microbial populations in their habitat (Alfreider et al., 1996; Llobet-Brossa et al., 1998; Murray et al., 1998; G16ckner et al., 1999; Simon et al., 1999). Several reviews discuss numerous aspects and applications of the method (Amann et al., 1995; 1997). In theory, each ribosome within a bacterial cell, containing one copy each of 5S, 16S and 23S rRNA, is stained by one probe molecule during the hybridization procedure, the high numbers of ribosomes per cell thus METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0 12-521530-4
All
Copyright © 2001 Academic Press Ltd rights of reproduction in any form reserved
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providing a natural signal amplification system (Figure ll.1).The method is mainly based on the rapidly increasing set of bacterial small subunit (16S rRNA) rRNA sequences, which has been gathered during the last decade for the study of microbial phylogeny (Woese et al., 1990; Ludwig and Schleifer, 1994). To a lesser extent, probes have been constructed that target the large subunit rRNA (23S rRNA) (Stoffels et al., 1998); this is, however, still hampered by the comparatively small number of available 23S rDNA sequences.
m.
Target
Probe
Figure 11.1. Schematic depiction of oligonucleotide binding.
FISH of bacteria was first described more than a decade ago (Giovannoni et al., 1988; DeLong et al., 1989; Amann et al., 1990b), and was hailed as a breakthrough for microbial ecology. However, researchers initially encountered discouraging difficulties when applying the method to environmental samples other than from highly eutrophic systems. The majority of bacteria in aquatic habitats is small, slowly growing or starving, and the signal intensities of hybridized bacterioplankton cells were frequently below detection limits or lost in high levels of background fluorescence. Accordingly, an early FISH protocol stated that there was '... a good deal of room for improvement of these techniques for practical field application.' (DeLong, 1993). This still holds true to a certain extent, but several important advances, in particular new quantitative protocols (G16ckner et al., 1996), brighter fluorochromes (Alfreider et al., 1996; G16ckner et al., 1996), commercial availability of probe labeling, advanced probe design software (Strunk et al., 1999) and better instrumentation have made the method attractive also for the less 'molecular' microbial ecologists.
FISH OF E N V I R O N M E N T A L SAMPLES O N M E M B R A N E FILTERS Principle Specificity of probe binding to the target site depends on the hybridization and washing conditions. Hybridization probes are added to a
208
defined, stringency-determining buffer at saturation concentrations (5 ng lJl ') to maximize probe binding. During hybridization the samples are incubated at elevated temperature in an airtight vessel saturated with water and formamide vapors of additional hybridization buffer to avoid concentration effects due to evaporoation. The washing step is performed at a slightly higher temperature and serves mainly to rinse off excess probe molecules at conditions that prevent unspecific binding. The protocols described focus on FISH with monolabeled fluorescent probes on membrane filters and glass slides. This approach is routinely used in our lab for the analysis of bacterioplankton and sediment samples of unknown composition (Llobet-Brossa et al., 1998; Pernthaler et al., 1998), and has been optimized for the processing of numerous samples. Other hybridization strategies (indirect probe labeling, signal amplification systems; Amann et al., 1992; Sch6nhuber et al., 1997) may be more appropriate for specific applications (e.g. FISH of autofluorescent cyanobacteria with horseradish peroxidase (HRP) labeled probes and Tyramide Signal Amplification (TSA); Sch6nhuber et al., 1999), but also need careful adaptation to the target microorganisms (e.g. cell wall permeabilization, fixation).
E q u i p m e n t and supplies General • • • • •
High quality epifluorescence microscope equipped with filter set for DAPI, FITC and CY3 Dry-type incubator or hybridization oven Water bath Freezer Fridge
Fixation of plankton samples
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• 100 ml glass bottles • Plastic Petri dishes (diameter, 5 cm) • White polycarbonate membrane filters (diameter, 47 mm; pore size, 0.2 IJm) • Cellulose nitrate support filters (diameter, 47 mm; pore size,_> 0.45 IJm) • Filter towers for 47 mm membrane filters • Vacuum pump • Particle-free 35% (w/v) formaldehyde solution (formalin) • 50, 80, and 96% (v/v) ethanol (only forTSA method)
Fixation and preparation of sediment samples • Microcentrifuge for 2 ml tubes • Vacuum pump • Ultrasonication probe • 2 ml screw-top microfuge tubes
209
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• 2 ml microfuge tubes • Plastic Petri dishes (diameter, 5 cm) • White polycarbonate membrane filters (diameter, 25 ram, pore size 0.2 pro) • Cellulose nitrate support filters (diameter, 25 ram, pore size _>0.45 lam) • Filter tower for 25 mm membrane filters • I× PBS • Ethanol • 4% (w/v) formaldehyde solution
Hybridization on filter sections and counterstaining • • • • • • • • • • • • • • •
•
2 ml microfuge tubes 0.5 ml microfuge tubes 50 ml polyethylene tubes and rack Blotting paper Razor blades Plastic Petri dishes Microscopic slides + cover slips CITIFLUOR mountant VECTA SHIELD mountant I MTris / HCI, pH 7.4 Formamide 0.5 M EDTA, pH 8 10% (w/v) sodium dodecyl sulfate (SDS) 5 M NaCI solution 4",6-Diamidino-2-phenylindole (DAPI) dissolved in distilled H~O, final concentration, I tJg ml' 80% (v/v) ethanol
Hybridization with horseradish peroxidase (HRP) labeled probes and T y r a m i d e Signal Amplification (TSA) • 2 ml microfuge tubes • 0.5 ml microfuge cubes • 50 ml polyethylene tubes and rack • Blotting paper • Razor blades • Plastic Petri dishes • Microscopic slides + cover slips • CITIFLUOR mountant • I MTris / HCI, pH 7.4 • Formamide • 0.5 M EDTA, pH 8 • 10% (w/v) sodium dodecyl sulfate (SDS) • 5 M NaCI solution • 10% blocking reagent (Roche) • TSA TM Fluorescence system (containing fluorophore labeled tyramide and amplification diluent) • TNT buffer (prepared according to the TSA TM Fluorescence system manual)
210
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Plate 2. Vital Stain and Probe (VSP) Technique. Micrographs derived from VSP triple staining of a surface water sample from the Barents Sea. (DAPI) conventional DAPI staining; (Probe) bacteria hybridized with FITC-labeled oligo nucleotide probes; and (PI) bacteria stained with propidium iodide.
Plate 3. Microautoradiography and probes. Micrograph of bacteria assayed by microautoradiography combined with fluorescence in situ hybridization (MICROFISH). (A) DAPI-stained bacteria (UV excitation). Dark spots surrounding cells are silver grains deposited on photographic emulsion around cells that took up tritiated free amino acids. (B) Bacteria hybridized with Cy3labeled oligonucleotide probe Eub 338 for eubacteria (green fluorescence). Cells with probe bound fluoresce yellow. Reprinted with permission from Cottrell and Kirchman (2000), Applied Environmental Microbiology.
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Plate 6. Satellite data collected at LEO-15 during July 16, 1999. (a) SeaWiFS estimated chlorophyll a for July 16, 1999 at Long-term Ecosystem Observatory (LEO15). Note that chlorophyll a values are highest, on average across the scene, within the cold waters observed within the AVHRR imagery. The black pixels to the northeast of the N line are pixels of 'bad' data, or areas which Equation (5) was not designed to accommodate. SeaWiFS chlorophyll a estimates in these areas are typically far above actual measurements. The region of black pixels is warmer than the colder upwelling regions, and are from a water mass being advected southward along the N e w Jersey coast. The origin of this water mass is u n k n o w n and illustrates some of the difficulties in using generalized oceanic relationships to describe coastal images. (b) An AVHRR image providing a m a p of sea surface temperature (varied by over 3°C). Arrow vectors of the sea surface currents (speed is given by the size of the arrow) as measured by a COastal RADar (CODAR) are shown to the southeast of Node A.
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Plate 7. Cross-shelf transects for (a) absorption (a(490) and a(550), m~) and (b) backscatter (bb(488) and bb(589), m ' ) on July 16, 1999 at the Long-term Ecosystem Observatory (LEO-15). The absorption data (at 490 and 550 nm) were collected with a Wetlabs absorption/attenuation meter (AC-9) that was factory calibrated prior to the cruise. Manufacturer-recommended protocols were employed to track AC-9 calibration by filtered air and double-distilled 0.2-micron-filtered water. Backscatter data (at 488 and 589 nm) was measured using a HOB1 Labs Hydroscat6. The optical instruments were mounted into a modified sea cage designed to minimize torsion that can impact instrument performance.
Assay: Fixation of plankton samples 1. Add formaldehyde to water sample to a final concentration of 2-4% and fix for at least i but no longer than 24 h. 2. A hand-operated vacuum pump and several autoclavable plastic filter towers that can be linked together for parallel sample processing make an inexpensive filtration apparatus for field work. Place the moistened support filter and polycarbonate filter into the filtration tower, and filter an appropriate volume of the fixed sample by applying gentle vacuum. Support filters may be utilized for several samples. For cell numbers of around 10'~ml ~, 10 ml of sample are sufficient. 3. After complete sample filtration, wash with 10-20 ml of sterile H~O; remove H,O by filtration. 4. Put membrane filter in Petri dish and allow to air-dry. 5. Store at -20°C until processing. Filters can be stored frozen for several months without apparent loss of hybridization signal.
Assay: Fixation and preparation of sediment samples 1. Suspend 0.5 ml of freshly collected sediment in 1.5 ml of 4% formaldehyde solution in a 2 ml screw-top microfuge tube. Fix for 1 to 24 h. 2. Centrifuge at 10 000 rpm for 5 min, pour off supernatant. 3. Add 1.5 ml of PBS and re-suspend sample. 4. Repeat steps 2 and 3. 5. Centrifuge at 10 000 rpm for 5 rain, pour off supernatant. 6. Add 1.5 ml of a 1:1 mix of PBS/ethanol and store sample at -20°C until further processing. 7. Re-suspend sample and transfer 20-100 lal of aliquot to 500 lJl of a 1:1 mix of PBS/ethanol in a 2 ml microfuge tube. 8. Sonicate aliquot for 20-30 s at low intensity using I s sonication pulses. If required, the sonicated sample can be further diluted. 9. Place cellulose nitrate support filters beneath the membrane filters to improve the distribution of cells. Add 15-20 M1 of aliquot from the sonicated sample to 2 ml of distilled water and filter this volume onto the membrane filters. 10. Air-dry filtered preparations and store in Petri dishes at -20°C until hybridization.
Assay: Fixation for the TSA method 1. Fix the water sample by adding the same volume of ethanol and store at -20°C until further treatment. Alternatively, freshwater samples may be immediately subjected to fixation and subsequent filtration onto membrane filters. 2. For the preparation of membrane filters, filter an aliquot (the volume depending on the density of the cells of interest; see above) of the fixed or fresh sample. 3. A second fixation (dehydration) is performed in the filtration apparatus by applying increasing concentrations of ethanol: cover the filter
211
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with 1 ml of 50% ethanol and incubate for 3 min, then remove liquid by filtration. This is repeated with 80% and 96% ethanol. . Filters are air dried and can be stored at -20 ° for several weeks.
Assay: H y b r i d i z a t i o n
of cells on m e m b r a n e
filters
1. Prepare 2 ml of hybridization buffer in a microfuge tube: Stock reagent
Volume
Final concentration in hybridization buffer
5 M NaC1 1 M Tris / HC1 Formamide Distilled H20 10% SDS (added last to avoid precipitation)
360 lal 40 pl % d e p e n d i n g on probe A d d to 2 ml 2 pl
900 mM 20 mM
0.01%
2. For the hybridization mixtures add 2 pl of probe working solution to 18 pl of hybridization buffer in a 0.5 ml microfuge tube; keep probe solutions dark and on ice. 3. Cut sections from m e m b r a n e filters with a razor blade. A 47 m m diameter filter should allow the preparation of 16-20 individual hybridizations. Label filter sections with a pencil, e.g., by n u m b e r i n g them. 4. Put filter sections on glass slides (upside facing up); several filter sections can be placed on one slide and hybridized simultaneously with the same probe. 5. Put a piece of blotting paper into a polyethylene tube and soak it with the remaining hybridization buffer. 6. A d d hybridization mix on the samples and place the slide with filter sections into the polyethylene tube (in a horizontal position). 7. Incubate at 46°C for at least 90 min (maximum: 3 h). 8. Prepare 50 ml of washing buffer in a polyethylene tube: Stock reagent
Volume
Final concentration in hybridization buffer
5 M NaCI
Depending on °7o formamide in hybridization buffer (Table 11.1)
900 mM
1 ml
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500 pl Add to 50 ml 50 pl
212
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Table I1.1 Concentrations of NaCI in washing buffer (48°C) at different concentrations of formamide in hybridization buffer (46°C) Formamide in hybridization buffer
washing buffer
(%)
(nnM)
0 5 10 15 20 25 30 35 40 45 50 55 60 65 70 75 80
900 636 450 318 225 159 112 80 56 40 28 20 14 10 7 5 3.5
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9. Quickly transfer filter sections into preheated washing buffer and incubate for 15 min at 48°C (water bath). 10. Pour washing buffer with filter sections into a Petri dish. Pick filter sections and rinse them by placing them into a Petri dish with distilled H20 for several seconds, then let them air-dry on blotting paper. 11. For counterstaining put filter sections on a glass plate, cover with 50 pl of DAPI solution, and incubate for 3 rain. The side holding bacterial cells should face up. Afterwards briefly rinse filter sections in distilled H~O, wash them for several seconds in 80% ethanol to remove unspecific DAPI staining, and air-dry. 12. Samples are m o u n t e d in a 4:1 mix of Citifluor and Vecta Shield. Vecta Shield contains a superior antibleaching reagent, but quenches DAPI fluorescence. The filter sections have to be completely dry before embedding, otherwise a fraction of cells will detach during inspection. 13. Double-stained and air-dried preparations as well as filters m o u n t e d on slides can be stored in the dark at -20°C for several days without substantial loss of probe fluorescence. 14. Probe-conferred fluorescence fades much more rapidly than DAP[ fluorescence in the microscopic image, and UV excitation will also bleach the CY3 signal. For counting, it is, therefore, safer to first quantify specifically stained cells in green excitation, and subsequently all 213
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cells from the same field of vision in UV excitation. At least 500 DAPIstained cells should be counted in plankton samples to obtain a counting error < +5% (Figure 11.2). 10 8 c-
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200
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,
400
00
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800
1000
1200
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cells (DAPI)
Figure 11.2. Effect of increasing cell counts on the precision of FISH results (plankton sample).
Assay: Hybridization with HRP labeled probes and T S A 1. The hybridization mixture described above is slightly changed by the replacing the 4001_ll of distilled H20 with the same volume of 10% blocking reagent (2% final concentration). This volume is reduced accordingly, if the concentration of formamide should exceed 60%. The 2~11 probe working solution should contain 100ng of HRPlabeled probe instead of fluorescently labeled oligonucleotides. 2. The next steps follow the assay of hybridization with fluorescently labeled probes except that the temperatures of hybridization and of washing are lowered to 35°C. This is required because of the instability of the enzyme at higher temperatures and is also reflected in different NaC1 concentrations for washing buffers (Table 11.2). 3. After washing and rinsing in distilled H20, transfer the filter sections directly to TNT buffer and equilibrate for 15 min at room temperature. 4. In the meantime, prepare substrate solution by mixing 1 volume of 40% dextrane sulfate in water, 1 volume of amplification diluent, and 1/50 volume of tyramide solution (the latter two are supplied with the TSA kit). About 20 lJl of the substrate solution is needed per filter section. 5. Remove excess liquid by briefly putting filters on blotting paper, but transfer them to a glass slide before they become completely dry. Cover them with substrate solution.
214
Table 11.2 Concentrations of NaCI in washing buffer (35°C) at different concentrations of formamide in hybridization buffer (35°C) Formamide in hybridization buffer
NaCI in washing buffer
(%)
(raM)
20 25 30 35 40 45 50 55 6O 65 70
145 105 74 52 37 26 19 13 9 8 5
6. Put the slide in another p o l y e t h y l e n e tube, the blotting p a p e r this time soaked by distilled water, and incubate for 30 to 90 min. 7. Rinse the filter pieces carefully in T N T buffer, i m m e r s e t h e m in the s a m e buffer and heat to 55°C for 15 min. 8. Rinse t h e m in distilled H~O and let them air-dry. 9. Cells can be counterstained with DAP! as described a b o v e or directly e m b e d d e d with Citifluor m o u n t a n t .
.~ .o
Troubleshooting •
Depending on the type of sediment, it might be necessary to also adapt the aliquot size prior to sonication. If too much sediment is suspended, sonication will lead to incomplete detachment of cells from particles. • Do not attempt to determine absolute cell counts from filters after hybridization, but only the percentage of hybridized cells.The distribution of cells on sections of a 47 mm diameter membrane filter is never as even as on a small filter, resulting in a higher error of the total DAPI counts. We find that, following our procedure, 80-90% of the initial bacterial cell numbers are recovered after hybridizations of bacterioplankton on membrane filters.This fraction may, however, depend on the type of sample and should be verified experimentally. • Hybridization stringency may also be adjusted by temperature rather than by the chemical compositon of buffers. We find that it is more convenient to keep incubator and water bath at one set temperature, so that several people can hybridize their samples in parallel irrespective of the probes they are using. • Due to the size of the HRP molecule, accessibility of probes to the cells may be discriminating.This is, for example, reflected in the preference for ethanol fixation rather than fixing with the crosslinking agents paraformaldehyde or
215
o'1" ,T
•
•
•
•
•
•
eeeeee
formaldehyde.The probability that not all organisms can be detected under the same conditions increases with the phylogenetic diversity of the target group. So it is recommended that the signal amplification method be used only for probes with a restricted target group for which fixation and hybridization conditions can be readily achieved. On white polycarbonate filters background fluorescence after DAPI staining is always somewhat worse than on black membrane filters. Black filters, however, show high levels of background fluorescence at green excitation. Use shorter DAPI staining time and/or longer ethanol washing to improve background. Make sure that hybridized filters have been thoroughly rinsed in distilled water before DAPI staining. Sometimes there can still be high or uneven background also in green excitation.To the untrained eye, a number of objects might appear to be hybridized bacteria, in particular autofluorescent debris (usually without DAPI signal) and cyanobacteria. Unhybridized filter sections should, therefore, be inspected sporadically to estimate the background level of'false positives'. Different fluorochromes are available for the TSA method. Among the kits one can choose between coumarin-, fluorescein-, tetramethylrhodamine, cyanine 3-, and cyanine 5-labeled tyramides. For the detection of cyanobacteria, fluorescein was superior to tetramethylrhodamine, as the difference between intrinsic autofluorescence and signal intensity was higher at blue excitation. For other applications different fluorescent dyes might be most suitable. It has been reported that a I ~ h pre-incubation of environmental samples with the antibiotic chloramphenicol (100 tJg ml' final concentration) in combination with an image intensifier equipment may result in significantly higher FISH detection rates in coastal marine samples (Ouverney and I=uhrman, 1997).We could not verify these results in our samples using standard epifluorescence microscopy. Prepare only small aliquots of probe working solutions (50-100 IJI). Use of repeatedly frozen and thawed probe working solutions may cause the appearance of numerous brightly fluorescent particles at green (CY3) excitation which do not show any signal in UV (DAPI) excitation. In addition, hybridization signals become dim and background is high. For HRP-labeled probes, do not freeze working solutions but store them in the fridge. Stock solution aliquots can be stored in the freezer, but once an aliquot is thawed to prepare working solutions, keep it at 4°C. Watch out for black cats and other signs of bad luck during hybridizations.
PROBE D E S I G N A N D T E S T I N G
Principle Presently, the major challenge of using FISH in environmental samples is no longer the staining technique itself, but rather the design and application of new probes within the context of a particular unknown microbial
216
assemblage. This approach, making use of FISH as but one in a suite of molecular biological techniques, has been termed 'full cycle rRNA analysis' (Amann et al., 1995). It refers to the initial construction of an rRNA clone library from environmental DNA after polymerase chain reaction (PCR) amplification with specific primers of 16S (23S) rDNA genes, comparative sequence analysis, subsequent probe design and the meaningful application of such new probes in the original habitat. Sequence information for probe construction may alternatively be obtained from bacteria isolated from the habitat by traditional means, e.g. by dilution techniques (Schut et al., 1993), but also from environmental RNA after reverse transcription into DNA or PCR re-amplification of excised individual bands obtained during denaturing gradient gel electrophoresis (DGGE) community fingerprinting (Rossell6-Mora et al., 1999). The melting behavior of oligonucleotide probes depends on temperature, the composition of the hybridization buffer, oligonucleotide sequence (G+C content), and on probe length. There are several strategies to determine stringent hybridization conditions for a newly designed probe, i.e. conditions that only allow hybridization of the probe to a perfectly matching template. They all rely on the experimental testing of the probe-target vs. probe-non-target stability. It would go beyond the scope of this chapter to provide detailed protocols for the various techniques used for probe testing, therefore here only one method based on image analysis (Neef et al., 1996) is described.
Equipment Appropriate software for probe construction, e.g. the program package ARB, released by the Technical University of Munich (Strunk et al., 1999). This program runs on different UNIX-based operating systems. It should work on a large database of currently > I 0 000 aligned 16S r D N A sequences, and additional sequences can easily be imported from various sources. The major drawback of ARB is a rather minimalistic online help system. The 'Probe Design Tool' of ARB allows the search for specific oligonucleotides of userdefined length and G+C content in a set of selected sequences, and to match these oligonucleotides with the ARB database. Potential probe candidates can then be BLASTed to GENBANK (http://www.ncbi.nlm.nih.gov/BLAST/) in order to check them against all currently available sequences. Teflon-coated multi-well slides. Mix of a 0.01% Cr K(SO4)2 / 0. 1% gelatin solution. Reagents and equipment for FISH (see above). Image analysis system: light-sensitive video or slow-scan CCD camera mounted on an epifiuorescence microscope, linked to a PC or Macintosh computer; image analysis software, e.g. the freely distributed program NIH
Image. (Alternatively: flow cytometer.) (Alternatively: reagents and equipment for membrane hybridizations.)
217
o'v i
It.
Probe design and synthesis 1. Preferably, full 16S rDNA sequences should be available when con-
2.
3.
4.
5.
structing specific probes. Partial sequences greatly limit probe design by reducing the number of potential target regions. In addition, partial sequences allow no decision about the specificity of existing probes that target unsequenced regions of the respective rRNAs. Not all sites within the ribosome are equally accessible for FISH, but may be blocked, e.g. by rRNA quarternary structure. The accessibility of the 16S rDNA of E. coli for oligonucleotide probes has recently been mapped by quantitative hybridization and flow cytometry (Fuchs et al., 1998). This color-coded map and a list of normalized hybridization intensities can probably not be directly transferred to other organisms, but due to the high evolutionary conservation of the ribosome it should give hints for probe design, and probe target sites promising high signal intensities can be selected. Ideally, a probe should show perfect base matching only with the target sequence(s), and have more than one base mismatch with the homologous region of non-target microorganisms. In order to minimize hybridization to such sites the base mismatches between probe and non-target rRNA should not be situated at the 3'- or 5'-ends of the oligonucleotide, but rather at a more central position. In addition, mismatches must be weighted according to their probability of undergoing non-Watson-Crick pairing with the base at the respective target position (e.g. a G-T mismatch is weaker than a G-A mismatch). The G+C content of an oligonucleotide influences duplex stability and therefore its melting behavior. Usually it should range between 50 and 60% for 18-24-base oligonucleotides. If the G+C content is too high or the probe is too long, stringent hybridizations may no longer be obtainable at our suggested hybridization temperature (see above). Readily labeled probes can be purchased from various companies. The indocarbocyanine fluorescent dye CY3, linked to the 5'-end of the oligonucleotide, to our knowledge is the brightest commercially available, routinely used label for FISH probes. An epifluorescence filter set specifically designed for CY3 excitation should be used in combination with the dye for optimal results.
Principles of probe testing 1. Probe stock are frequently delivered lyophilized. Suspend in 100 lal sterile H20. To determine probe concentration measure the absorbance of a 1:100 diluted stock solution at 260 nm (1 Ae,,,,~ 20 lag ml ' DNA). This information is usually supplied by the manufacturer, but it is still useful to perform this measurement in order to check the quality of the labeling. The dye CY3 shows maximum absorbance around 550 nm, and the ratio A,,.JAs-,,~ should be approximately 1 for a monolabeled
218
18-mer. Aliquots of working solutions are prepared at concentrations of 50 ng pl ~and stored in the dark at -20°C. Lyophilized HRP-labeled probes are suspended in sterile H20, too. For calculating the concentration it has to be taken into account that the enzyme itself contributes to the measured absorbance at 260 nm. Therefore the measured A ..... has to be lowered by k x A .... with the correction factor k = 0.276. Presuming optimal labeling the ratio of the measured A:,,, and A4,,~ should be around 3. Aliquots of the stock solution can be stored in at -20°C, working solutions prepared at concentrations of 50 ng pl' should be stored at 4°C (see troubleshooting on repeated freezing and thawing of working solutions). 2. The 'gold standard' of testing new probes for FISH is hybridization of isolates that show no and one mismatch with the oligonucleotide, respectively, at increasing levels of stringency. The simplest way of establishing stringency is by changing the concentrations of formamide in the hybridization buffer (in steps of 5%) at one fixed incubation temperature rather than by changing hybridization temperature. Change of fluorescence intensities of individual cells can then be quantified, e.g. by computer-assisted image analysis (Neef et al., 1996) or by flow cytometry (Fuchs et al., 1998). Adequate hybridization stringency is often the highest concentration of formamide in the hybridization buffer that does not result in loss of fluorescence intensity of the target cells. At this concentration, hybridization to the nontarget organism should no longer occur. As a rule of thumb, an 18-base oligonucleotide with a G+C content between 50-60% will start to dissociate at a concentration of approximately 30-40~ of formamide in the hybridization buffer when hybridized in our buffer at 46°C. Alternatively, probes can be radiolabeled and hybridized to extracted rRNAs that have been blotted onto nylon membranes (Stahl and Amann, 1991). 3. Frequently new probes will be designed to target bacterial groups that are only known from their rDNA sequences. In this case it is not possible to test these probes on isolates. Two strategies are available to confirm probe specificity: (i) the rDNA to be tested can be transcribed in vitro into RNA, which is then blotted on a nylon membrane and hybridized to the radiolabeled oligonucleotide at increasing levels of formamide (e.g. Pernthaler et al., 1998); (ii) FISH with the new probe is carried out directly on environmental samples. The relative abundance of detected cells and the average cell brightnesses are determined at different formamide concentrations by counting and image analysis. At stringency levels that are too low, cell counts may be higher because non-target populations are also detected. If stringency is too high, mean cell brightness of the target population will rapidly decline (Figure 11.3). 4. To quickly evaluate the brightness of a probe and thus its sensitivity in environmental samples it may be helpful to hybridize bacteria from a stationary phase culture of the target organism and compare signal intensities with those of a hybridization with a general bacterial or universal probe. 219
,u
i U.
100 E-80
co
I
I
I
I
60
03 ctO')
0 c-
N
40
20
0
I
10
I
20
I
I
30
40
I
I
50
% Formamide
Figure 11.3. Melting curve equivalent of a probe targeted to an uncultivated freshwater bacterial phylotype, determined from hybridizations of environmental samples at increasing levels of formamide and subsequent image analysis (mean _+SE).
Assay: FISH of pure cultures on multi-well glass slides for probe testing 1. Heat the Cr K(SO~):/gelatin solution to 65°C, soak multi-well slides for 2 s and air-dry them. 2. Harvest cells during logarithmic growth, fix aliquot with formalin (2% final concentration) for 1-24 h. 3. Centrifuge cells (at 14 000 rpm). 4. Pour off supernatant, add 1 ml of PBS and resuspend cells. 5. Repeat steps 3 and 4. 6. Centrifuge cells (at 14 000 rpm), pour off supernatant, add 1 ml of a 1:1 mix of PBS/ethanol and resuspend cells; at this stage samples can be stored at -20°C for several months. 7. Spot 2-20 pl of fixed cell suspension on gelatin-coated slide and airdry. 8. Prepare hybridization buffers with different formamide concentrations as described above. 9. Prepare hybridization vessels from polyethylene tubes as described above; use separate tubes for each concentration of formamide; the buffer reservoir in each air-tight vessel must be the correct hybridization buffer. 10. For each hybridization (well) mix 1 pl of probe working solution and 9 pl of hybridization buffer. 11. Hybridize and wash as described.
220
Assay: Cell brightness quantification by image analysis 1. Hybridize fixed cells on multi-well slides at different concentrations of formamide. 2. Capture gray scale images in all samples at one fixed exposure condition; depicted cells must contain no overexposed pixels. 3. Pre-filter gray scale image to enhance cell edges, e.g. by 'Unsharp Masking': subtract a repeatedly low-pass filtered ('blurred') image from the original and subsequently multiply the resulting image by an appropriate factor (10-20). 4. Threshold the processed gray scale image to produce a binarized image containing only objects and background 5. Set lower and upper levels of accepted object size in the binarized image to eliminate pixel noise and clustered cells. 6. Determine the cell brightnesses of all objects in the mask image by overlaying it with the original gray scale image. Average the mean object brightnesses of several images (5-10) to determine means and standard deviations per image.
Troubleshooting • To increase the specificity of probes that show only one mismatch with nontarget sequences, it may be required to construct one or several competitor oligonucleotides. Such competitors are designed to perfectly match with the non-target sequence at the homologous site, and they are subsequently synthesized without fluorescent label. During hybridization, the competitor is added to the buffer at an equal concentration as the labeled probe. This strategy has been successfully applied to construct a pair of 23S rRNAtargeted probes that discriminate between the beta and gamma subclasses of the Proteobocterio (Manz et o1., 1992). • One may encounter unresolved bases (N, R,¥, K .... ) rather than mismatches at the target region of database sequences from non-target organisms. In this case, it is impossible to know if such sequences will be detected by the probe. However, after careful comparison with closely related sequences some intelligent guesses about the likelihood of accidental base identity may be possible (e.g. is the position conserved or highly variable etc.). If possible, check probe specificity experimentally. • Sometimes we observe that FISH signal increases when adding some formamide to the hybridization buffer.We assume that formamide helps to enhance probe accessibility.Therefore during initial testing probes should be used with at least 10% of formamide in the hybridization buffer. • A 'quick and dirty' way to decide if newly designed probes are working at all (i.e. if the target site is, in principle, accessible) is to perform FISH of environmental samples at increasing levels of formamide followed by the counting of hybridized cells at different stringencies and a subjective estimate of signal intensity (e.g. in three arbitrary brightness classes). • Frequently one cannot find perfect probes. Probe design represents a search for compromise of advantages (e.g. full-group coverage) and drawbacks (e.g. outside-group hits). One should also bear in mind that 15 to 30-base oligo-
221
m U.
nucleotide probes are as reliable as the database from which they are constructed.The influx of new rRNA sequence information to the databases may eventually require a re-evaluation of probe specificity and even redesign of probes that can no longer be regarded as specific. For the reliable detection of bacterial lineages or phylotypes without cultivated representatives, it is, therefore, advantageous to construct at least two parallel probes to different positions of the target 16S rRNA(s).
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APPLICATIONS During recent years FISH has been successfully applied in freshwater, coastal and offshore marine planktonic habitats, as well as in coastal sediments. It has been shown that the fraction of bacteria detectable by FISH corresponds well with the abundance of active cells as determined by microautoradiography in coastal marine bacterioplankton (Karner and Fuhrman, 1997). Group-specific probes for different subclasses of the Proteobacteria have, for example, been utilized to study the composition of lake snow (Weiss et al., 1996) and of the microbial assemblages in the plankton and winter cover of a mountain lake (Alfreider et al., 1996), to demonstrate fundamental differences in the composition of freshwater and marine bacterioplankton (G16ckner et al., 1999), or to show vertical zonations of the microbial communities in m u d d y and sandy Wadden Sea sediments (Llobet-Brossa et al., 1998). The seasonal dynamics of members of major phylogenetic lineages and of several individual bacterial phylotypes have been studied in lake bacterioplankton (Pernthaler et al., 1998). Blooms of members of the Cytophaga/Flavobacterium cluster have been described at the Antarctic marginal ice zone during a Phaeocystis bloom (Simon et al., 1999). Archaeal seasonal abundances and vertical distributions in Antarctic coastal waters were followed by FISH with oligonucleotide and polynucleotide probes (Murray et al., 1998; DeLong et al., 1999). FISH was also applied in laboratory and field studies to prove that protistan predation can shift community composition in freshwater bacterioplanktonic assemblages (Pernthaler et al., 1997b; gimek et aI., 1997; gimek et al., 1999). Other applications of FISH with rRNA-targeted oligonucleotides in, for example, soil or wastewater treatment systems, are too numerous to be listed here. An overview of early applications can be found in Amann et al. (1995).
e e e e e , e, C O N C L U S I O N S The low fluorescence intensities of hybridized aquatic bacteria still represent a partially unresolved problem of the method presented here. Due to low signal levels, it is presently not feasible to use flow cytometry, the ideal tool for directly counting FISH-stained cells, in plankton samples. Microscopically, a variable fraction of cells detected by general nucleic 222
acid staining (e.g., DAPI) are also visualized by FISH with more general probes. We find that, d e p e n d i n g on the season, 30% to more than 80% of DAPI-stained objects in the G e r m a n Bay hybridize with the general bacterial probe EUB338 (Pernthaler et al., unpublished data). D e p e n d i n g on the philosophy of the researcher, the FISH-undetectable fraction has been dismissed as dead 'ghosts' (Zweifel and Hagstr6m, 1995), or classified as living, non-growing bacteria. Relatively new groups of bacteria that were only recently found to be a b u n d a n t in some environments by cultivationi n d e p e n d e n t methods, are not detected by the bacterial probe EUB338 (Daims et al., 1999). Moreover, there is ample evidence that a fraction of the cells not hybridizing with a general bacterial probe (EUB338) are members of the archaea (DeLong et al., 1994; F u h r m a n and Ouverney, 1998). We believe that eventually even presently undetectable cells will be detected with i m p r o v e d FISH techniques: multiple-labeled polynucleotide probes have been described (Trebesius et al., 1994; DeLong et al., 1999) that promise to overcome the problem of low signal intensity in oligotrophic open ocean environments. However, the synthesis and testing of such probes is much more technically d e m a n d i n g than FISH with oligonucleotide probes.
References Alfreider, A., Pernthaler, J., Amann, R., Sattler, B., Gl6ckner, F.-O., Wille, A. and Psenner, R. (1996). Community analysis of the bacterial assemblages in the winter cover and pelagic layers of a high mountain lake by in situ hybridization. Appl. Envitvn. Microbiol. 62, 2138 2144. Amann, R., Snaidr, J., Wagner, M., Ludwig, W. and Schleifer, K.-H. (1996). In situ visualization of high genetic diversity in a natural microbial community. J. Bacteriol. 178, 3496-3500. Amann, R., Gl6ckner, E O. and Neef, A. (1997). Modern methods in subsurface microbiology iH situ identification of microorganisms with nucleic acid probes. FEMS Microbiol. Rev. 20, 191-200. Amann, R. l., Binder, B. J., Olson, R. J., Chisholm, S. W., Devereux, R. and Stahl, D. A. (1990a). Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Appl. Environ. Microbiol. 56, 1919-1925. Amann, R. I., Krumholz, L. and Stahl, D. A. (1990b). Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J. Bacteriol. 172, 762-770. Amann, R. 1., Zarda, B., Stahl, D. A. and Schleifer, K. H. (1992). Identification of individual prokaryotic cells by using enzyme-labeled, rRNA-targeted oligonucleotide probes. Appl. Environ. Microbiol. 58, 3007-11. Amann, R. 1., Ludwig, W. and Schleifer, K. H. (1995). Phylogenetic identification and i17 situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59, 143-169. Daims, H., Bruhl, A., Amann, R., Schleifer, K. H. and Wagner, M. (1999). The domain-specific probe EUB338 is insufficient for the detection of all Bacteria: Development and evaluation of a more comprehensive probe set. Syst. Appl. Microbiol. 22, 434-444. DeLong, E. E, Wickham, G. S. and Pace, N. R. (1989). Phylogenetic stains: ribosomal RNA-based probes for the identification of single cells. Science 243, 1360-1363. 223
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o3: I.L
DeLong, E. E (1993). Single-cell identification using fluorescently labeled ribosomal RNA-specific probes. In: Handbook of Methods in Aquatic Microbial Ecology (P. Kemp, B. E Sherr, E. B. Sherr and J. Cole, Eds), pp. 285-294. Lewis Publishers, Boca Raton. DeLong, E. E, Wu, K. Y., Prezelin, B. B. and Jovine, R. V. M. (1994). High abundance of Archaea in Antarctic marine picoplankton. Naturr 371, 695-697. DeLong, E. F., Taylor, L. T., Marsh, T. L. and Preston, C. M. (1999). Visualization and enumeration of marine planktonic archaea and bacteria by using polyribonucleotide probes and fluorescent in situ hybridization. Appl. Environ. Microbiol. 65, 5554-5563. Fuchs, B. M., Wallner, G., Beisker, W., Schwippl, I., Ludwig, W. and Amann, R. (1998). Flow cytometric analysis of the in-situ accessibility of Escherichia coli 16S rRNA for fluorescently labeled oligonucleotide probes. Appl. Environ. Microbiol. 64, 4973-4982. Fuhrman, J. A. and Ouverney, C. C. (1998). Marine microbial diversity studied via 16S rRNA sequences: cloning results from coastal waters and counting of native Archaea with fluorescent single cell probes. Aquatic Ecol. 32, 3-15. Giovannoni, S. J., DeLong, E. E, Olsen, G. J. and Pace, N. R. (1988). Phylogenetic group-specific oligodeoxynucleotide probes for identification of single microbial cells. J. Bacteriol. 170, 720-726. GI6ckner, E O., Amann, R., Alfreider, A., Pernthaler, J., Psenner, R., Trebesius, K. and Schleifer, K.-H. (1996). An iu situ hybridization protocol for detection and identification of planktonic bacteria. Syst. Appl. Microbiol. 19, 403-406. GI6ckner, E O., Fuchs, B. M. and Amann, R. (1999). Bacterioplankton compositions of lakes and oceans: a first comparison based on fluorescence in situ hybridization. Appl. EnviJvn. Microbiol. 65, 3721-3726. Karner, M. and Fuhrman, J. A. (1997). Determination of active marine bacterioplankton: A comparison of universal 16S rRNA probes, autoradiography, and nucleoid staining. Appl. Envirolz. Microbiol. 63, 1208-1213. Llobet-Brossa, E., Rossell6-Mora, R. and Amann, R. (1998). Microbial community composition of Wadden sea sediments as revealed by fluorescence in situ hybridization. Appl. EnviroJz. Microbiol. 64, 2691-2696. Ludwig, W. and Schleifer, K.-H. (1994). Bacterial phylogeny based on 16S and 23S rRNA sequence analysis. FEMS Microbiol. Rev. 15, 155-173. Manz, W., Amaim, R., Ludwig, W., Wagner, M. and Schleifer, K.-H. (1992). Phylogenetic oligodeoxynucleotide probes for the major subclasses of Proteobacteria: Problems and solutions. Syst. Appl. Microbiol. 15, 593-600. Murray, A. E., Preston, C. M., Massana, R., Taylor, L. T., Blakis, A., Wu, K. and Delong, E. E (1998). Seasonal and spatial variability of bacterial and archaeal assemblages in the coastal waters near Anvers Island, Antarctica. Appl. Envirosl. Microbiol. 64, 2585-2595. Neef, A., Zaglauer, A., Meier, H., Amann, R., Lemmer, H. and Schleifer, K. H. (1996). Population analysis in a denitrifying sand filter: conventional and in situ identification of Paracoccus spp. in methanol-fed biofilms. Appl. Environ. Microbiol. 62, 4329-39. Ouverney, C. C. and Fuhrman, J. A. (1997). Increase in fluorescence intensity of 16S rRNA in situ hybridization in natural samples treated with chloramphenicol. Appl. Environ. Microbiol. 63, 2735-2740. Pernthaler, J., Posch, T., Simek, K., Vrba, J., Amann, R. and Psenner, R. (1997). Contrasting bacterial strategies to coexist with a flagellate predator in an experimentaI microbial assemblage. Appl. Environ. Microbiol. 63, 596-601. Pernthaler, J., GlOckner, E O., Unterholzner, S., Alfreider, A., Psenner, R. and Amann, R. (1998). Seasonal community and population dynamics of pelagic
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Bacteria and Archaea in a high mountain lake. Appl. Environ. Microbiol. 64, 4299-4306. Rossell6-Mora, R., Thamdrup, B., SchSfer, H., Weller, R. and Amann, R. (1999). The response of the microbial community of marine sediments to organic input under anaerobic conditions. Syst. Appl. Microbiol. 22, 237-248. Sch6nhuber, W., Fuchs, B., Juretschko, S. and Amann, R. (1997). Improved sensitivity of whole-cell hybridization by the combination of horseradish peroxidaselabeled oligonucleotides and tyramide signal anaplification. Appl. Environ. Microbiol. 63, 3268-3273. Sch6nhuber, W., Zarda, B., Eix, S., Rippka, R., Herdman, M., Ludwig, W. and Amann, R. (1999). In situ identification of cyanobacteria with horseradish peroxidase-labeled, rRNA-targeted oligonucleotide probes. Appl. Environ. Microbiol. 65, 1259-1267. Schut, E, De Vries, E. J., Gottschal, J. C., Robertson, B. R., Harder, W., Prins, R. A. and Button, D. K. (1993). Isolation of typical marine bacteria by dilution culture: growth, maintenance, and characteristics of isolates under laboratory conditions. Appl. Environ. Microbiol. 59, 2150-2160. Simek, K., Vrba, J., Pernthaler, J., Posch, T., Hartman, P., Nedoma, J. and Psenner, R. (1997). Morphological and compositional shifts in an experimental bacterial community influenced by protists with contrasting feeding modes. Appl. Environ. Microbiol. 63, 587-595. Simek, K., Kojecka, P., Nedoma, J., Hartman, P., Vrba, J., and Dolan, J. R. (1999). Shifts in bacterial community composition associated with different microzooplankton size fractions in a eutrophic reservoir. Liranol. Oceanogr. 44, 1634-1644. Simon, M., G16ckner Frank, O. and Amann, R. (1999). Different community structure and temperature optima of heterotrophic picoplankton in various regions of the Southern Ocean. Aquatic Microbial Ecol. 18, 275-284. Stahl, D. A. and Amann, R. (1991). Development and application of nucleic acid probes. In: Nucleic Acid Teclmiques in Bacterial Systematics (E. Stackebrandt and M. Goodfellow, Eds), pp. 205-248. John Wiley and Sons Ltd., Chichester, UK. Stoffels, M., Amann, R., Ludwig, W., Hekmat, D. and Schleifer, K.-H. (1998). Bacterial community dynamics during start-up of a trickle-bed bioreactor degrading aromatic compounds. Appl. Environ. Microbiol. 64, 930-939. Strunk, O., Gross, O., Reichel, B., May, M., Hermann, S., Stuckmann, N., Nonhoff, B., Ginhart, T., Vilbig, A., Lenke, M., Ludwig, T., Bode, A., Schleifer, K.-H. and Ludwig, W. (1999). ARB: a software environment for sequence data. Department of Microbiology, Technische Universitfit Mflnchen, Munich, Germany. Trebesius, K., Amann, R., Ludwig, W., Mtihlegger, K. and Schleifer, K.-H. (1994). Identification of whole fixed bacterial cells with nonradioactive 23S rRNAtargeted polynucleotide probes. Appl. Environ. Microbiol. 60, 3228-3235. Wagner, M., Schmid, M., Juretschko, S., Trebesius, K. H., Bubert, A., Goebel, W. and Schleifer, K. H. (1998). In situ detection of a virulence factor mRNA and 16s rRNA in listeria monocytogenes. FEMS Microbiol. Lett. 160, 159-168. Weiss, P., Schweitzer, B., Amann, R. and Simon, M. (1996). Identification in situ and dynamics of bacteria on limnetic organic aggregates (lake snow). Appl. Environ. Microbiol. 62, 1998-2005. Woese, C. R., Kandler, O. and Wheelis, M. L. (1990). Towards a natural system of organisms: proposal for the domains Archaea Bacteria and Eucarya. Proc. Natl. Acad. Sci. USA 87, 4576-4579. Zweifel, U. L. and Hagstr6m, A. (1995). Total counts of marine Bacteria include a large fraction of non-nucleoid-containing Bacteria (Ghosts). Appl. E~;viro;;. Microbiol. 61, 2180-2185.
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List of suppliers The following is a selection of companies. For most products, alternate suppliers can be found. ARB http://www.mikm.biologie.tumuenchen.de/pub/ARB/
INTERACTIVA The Virtual Laboratory Sedanstr. 10 89077 Uhn Germany Tel.: +49 731 / 93579-290 Fax: +49 731 / 93579-291 http: //www.interactiva.de/
Probe design software Carl Zeiss KiYnix,sallee 9-21 D-37081 GiJttingen Germany Teh +49 1803 33 63 34 Fax: +49 551 5060 480 h t tp :/ /www.zeiss.de/
Probe synthesis and labeling (Cy3 and HRP-labeling) Nalge Nunc International 2000 North Aurora Road Naperville, IL 60563, USA Tel: +1 630 983-5700 Fax: +1 630 416-2519 http://www.nalgenunc.com
High-end epifluorescence microscopes Chroma Technology Corp. 72 Cotton Mill Hill, Unit A-9 Brattleboro, VT 05301, USA Teh +1 802-257-1800 Fax +1 802-257-9400 http://www.chroma.com/
Plastic filtration towers, hand pump
High quality filter set for Cy3 Citifluor Ltd 18 Enfield Cloisters Fanshaw Street London N1 6LD, UK Fax: +44 01 227 827724
NEN® Life Science Products, Inc. 549 Albany Street Boston, M A 02118, USA Tel.: +1 800-551-2121 Fax: +1 617-482-1380 h t tp :/ /www.nen l ifesci.com /
TSA T M Fluorescence Systems NIH Image http://rsb.info.nih.gov/nih-image/
Citifluor AF1 mountant Diagnostic Instruments, Inc. 6540 Burlvughs Street Sterling Heights, MI, USA Teh +1 810 731-6000 Fax: +1 810 731-6469 http://wwz~.diaginc.com/index.htm
Digital cameras
Image analysis software Vector Laboratories, Inc. 30 Ingold Road Burlingame, CA 94010, USA Tel.: +1 650 697-3600 Fax: +1 650 697-0339 http://zL,ww.vectorlabs.com/
VectaShield mountant
226
12 Measuring Bacterial Biomass Production and Growth Rates from Leucine Incorporation in Natural Aquatic Environments David Kirchman College o( Marine Studies, University of Deloware, Lewes,DE 19958, USA
CONTENTS
Introduction Measuring3H-leucine incorporation by the filter method Measuring3H-leucine incorporation by the microcentrifuge method Conclusions
~,~I,~,~,4,~, I N T R O D U C T I O N The rate of biomass production is a fundamental property of all organisms in nature, but it is an especially important parameter of microbes in natural aquatic environments. An estimate of microbial production can be used as a general index of microbial activity and specifically to calculate growth rates. Since m a n y processes scale with it, biomass production can be used to obtain a first-order estimate of rates of several processes mediated by microbes. For example, in the case of heterotrophic bacteria, the organisms to be discussed here, biomass production can be used to estimate use of dissolved organic material (DOM) if coupled with an estimate of the growth efficiency. That is, DOM uptake equals bacterial biomass production divided by the growth efficiency (expressed as a fraction, not a percentage). Biomass production is the increase in biomass per unit time per unit w)lume or per area and is a function of both biomass (B), usually expressed as carbon mass per volume (e.g. lagC l~), and the specific growth rate (ia) (e.g. h ') (Ducklow, 2000). in the absence of any mortality (grazers and viruses), bacterial biomass increases exponentially (although not necessarily very quickly) which can be described by dB/dt METt tODS IN MICROBIOLOGY, VOLUME 30 ISBN 0 12 521530-4
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(12.1)
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In this case, bacterial production is the first derivative ( d B / d t ) or the slope of the curve on graphs of biomass (B) versus (t). If the data are graphed as ln(B) versus time, then the slope of this semi-log graph is the growth rate (V). In nature, bacteria seldom if ever live in the absence of protist grazers and viruses, both of which cause bacterial mortality. In most natural systems, most of the time, bacterial production is matched by mortality such that (dB/dt),,~.~ = 0. However, the methods discussed here measure 'gross production', i.e. biomass production unaffected by mortality. It is the rate of biomass production that would occur if mortality were zero. This is possible because the methods rely on incubations that are relatively short (hour or less) compared to the timescale of bacterial growth and mortality (a day or longer). Although we say we measure 'gross production', the rate does not include respiration. Ducklow (2000) discusses the differences in uses of the terms 'gross' and 'net' production by bacterial and phytoplankton ecologists. Ducklow (2000) also reviews the m a n y methods that have been used to examine bacterial production in aquatic environments. Although alternative methods are still valuable for specific applications, recent investigations of natural aquatic environments have used either thymidine (TdR) incorporation or leucine (Leu) incorporation or both to estimate bacterial production. The two methods, which were originally proposed by Fuhrman and Azam (1980) and Kirchman et al. (1985), respectively, have many parallels, and the experimental details are nearly identical. Both are rapid, easy, and specific for heterotrophic bacteria, some of the main reasons w h y they have been widely adapted by microbial ecologists. I focus here on the leucine method because it is more straightforward than the TdR method to estimate bacterial production; if one relies on 'theoretical' conversion factors, the only variable assumption in extrapolating from Leu incorporation to bacterial production is the degree of isotope dilution (Kirchman, 1993), as described below. Consequently, over the last few years more investigators seem to be using Leu incorporation rather than the TdR method.
M E A S U R I N G 3H-LEUCINE I N C O R P O R A T I O N B Y T H E FILTER M E T H O D
Principle Two variations of the Leu method will be described, but the basic biochemistry and physiology behind the methods are the same. Leucine incorporation is often used to measure protein synthesis in pure cultures of bacteria because leucine is a constant proportion of all protein (7.3~7~; Kirchman at al., 1985). Consequently, rates of protein synthesis can be estimated from the rate at which this amino acid appears in the protein fraction. The original method called for hot trichloroacetic acid (TCA) extraction to measure leucine incorporated specifically into protein, but this step is not necessary because the difference between a cold and hot 228
TCA extraction is negligible; nearly all leucine is incorporated directly into protein (Kirchman et al., 1985). Rates of total biomass production can be estimated in turn if the a m o u n t of protein per cell or per cellular mass is known. Although cell size and thus protein per celt can vary greatly, protein is a relatively constant fraction of bacterial biomass (60% of dry weight; Simon and Azam, 1989). Incorporation of leucine (and thymidine) is dominated by heterotrophic bacteria in most aquatic habitats. Rates of leucine incorporation into protein are estimated from the appearance of radioactivity, added as 'H-leucine, in the protein fraction. The added ~H-leucine (20 nM) normally is much higher than in situ concentrations (< 1 nM). This high added concentration has two effects. First, it means that the natural extracellular leucine usually can be ignored in all calculations. Second, because extracellular concentrations are so high with the added leucine, bacteria will take up the exogenous ~H-leucine and repress leucine biosynthesis, i.e. the production of non-radioactive leucine which is subsequently used for protein synthesis. But the problem is, some a m o u n t of leucine biosynthesis, usually unknown, may continue even in the presence of high exogenous leucine, thus 'diluting' incorporation of radioactive leucine. This is called 'isotope dilution' (ID). In summary, the equation describing biomass production as estimated from leucine incorporation (Leu incorp) is: Biomass production = Leu Incorp x 131.2 + (Leu per protein) x (cell C per protein) x ID (12.2) which gives biomass production as gC per volume per unit time of the incubation. The molecular weight of leucine is 131.2 and converts moles of leucine incorporation into grams of C. The fraction of Leu in protein ('Leu per protein') is 7.3% (0.073 in the equation) and cellular C per protein is 0.86 (Simon and Azam, 1989). The few measured estimates of ID average to about 2 (Simon and Azam, 1989). A more conservative approach (lower production) would assume that ID = 1. In which case, bacterial production can be calculated as: Biomass production = Leu lncorp x 1.5 kg C per mol
(12.3)
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when leucine incorporation is measured in moles incorporated per unit time and volume. Specific growth rates can be estimated from rates of biomass production if a few assumptions are made. If we apply Equation (12.1) to gross production (not net production), i.e. that measured by the TdR or Leu approaches, then the specific growth rate (~1) is: p - BP/B
(12.4)
where B P is biomass production (biomass per time unit per volume or area) and B is a measure of bacterial abundance (cells per volume) or biomass (cellular mass per volume). Cellular biomass is usually estimated by multiplying bacterial abundance by a carbon per cell conversion factor, 229
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c
e.g. 10 fgC per cell (Fukuda et al., 1998). Biomass versus cell production is discussed below. Note that growth rates have units of 'per time', e.g. d 1, whereas generation time (g) is g = ln(2)/p = 0.693/p
(12.5)
and has units of 'time', e.g. hours or days. Ecologists often discuss 'doublings per day' which has the same units as specific growth rates, but is calculated as 1/g. The growth rate calculated from biomass production would reflect an average for the entire bacterial assemblage, including both the very slow and the very fast growing cells. The extreme is the case of nonviable cells with a growth rate of zero. Over the years, there has been much debate about the fraction of inactive (if not dead) bacterial cells in aquatic habitats, but generally a high proportion of cells do incorporate leucine (Kirchman et al., 1985) and thymidine (Fuhrman and Azam, 1982). However, regardless of the capacity of bacteria to incorporate these two compounds, several studies do suggest that activity and presumably growth rates vary greatly among various members of the bacterial assemblages (del Giorgio et al., 1997; Sherr et al., 1999). Although the limitations are obvious, growth rates calculated by the method outlined above can be very useful in characterizing bacterial assemblages. Also, there are few alternative approaches.
Equipment and reagents • Tritiated leucine ([2,3,4-3H] leucine) as a stock solution with specific activity of >60 nmol per Ci (I mCi ml -~) (NEN,Amersham). • Polycarbonate incubation tubes or flasks of appropriate sizes for environment. Other materials (e.g. glass or other plastics) should be avoided. • Vacuum pump, flasks and filter holders (25 mm) for filtering radioactive and corrosive liquids. • Filters of mixed cellulose esters with pore sizes of 0.45 pm (or 0.22 lum) and diameter of 25 mm (Millipore).Two pairs of forceps. It may be necessary to remove the plastic handle so that the forceps can fit down to the bottom of a 7 ml scintillation vial. • Pipettes (e.g. Pipetman) that dispense volumes ranging from microliters (for the 3H-leucine) to milliliters. Repeating dispensers for the ethyl acetate and scintillation cocktail. • Trichloroacetic acid (TCA) in a stock solution of 50% and as a wash solution (5%) • Ethanol, 80%. • Ethyl acetate. • Scintillation cocktail and scintillation vials (7 ml). Ultima-Gold (Packard Instruments) was found to be the optimal cocktail (Ducklow, personal communication).
230
• • •
Vortexer. Scintillation counter (e.g. Beckman,Wallac). Appropriate containers for radioactive corrosive liquids and radioactive solids.
Assay
It is not necessary to use sterile techniques to conduct the following assay. Contamination by organic material, however, should be avoided as it could stimulate bacterial growth or dilute the added 3H-leucine. Plastic gloves should be worn at all times to protect the sample from contamination. 1. Add an appropriate volume of the water sample to an incubation vessel. The volume may range from 5 ml of highly active waters to 100 ml of very inactive waters. Duplicates or triplicates should be prepared per sample. 2. Prepare a killed control by adding TCA to a final concentration of 1%. Other killing agents can be used (e.g. formaldehyde), but TCA is needed for other steps in the assay anyway and it does not affect the incubation vessel. Bottles exposed to formaldehyde or gluteraldehyde should not be used for live incubations. Also, formaldehyde and gluteraldehyde need to be handled in a hood. 3. Add 3H-leucine to the samples and to the killed control and then mix by hand. The final concentration of leucine should be 20 riM, which usually is sufficient to maximize incorporation rates, indicating that isotope dilution has been minimized. This maximum concentration can be determined empirically by simply varying the added 3H-leucine concentration and noting at what concentration 3H-leucine incorporation reaches a maximum. The actual volume of ~H-leucine added per incubation will vary, depending on the incubation volume and the specific activity of the 3Hleucine batch from the manufacturer. 4. Incubate the samples at the in situ temperature for an appropriate time. The incubation time may vary from 30min to 24h, depending on the activity level. For many applications, end point determinations are quite adequate, but for unknown environments it is advisable to measure incorporation over time in order to determine the best single incubation time. The best time is the shortest period that gives a measurable rate with acceptable errors. For most water samples, this time will be about one hour. Water samples without visible particles probably do not need to be shaken during the incubation, but if material is present that may settle, it is advisable to shake gently. 5. After incubation, filter samples through filters using minimal vacuum (<200 mm Hg). The start and end times of the incubation are noted. 6. With the vacuum connected, rinse the filter twice with 3 ml of icecold TCA.
231
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7. Rinse twice with 3 ml of ice-cold 80% ethanol. 8. With the vacuum still connected, lift the tower off the filter and use a small volume of ethanol to rinse the filter edge that was covered by the tower. 9. Using two forceps, fold the filter into quarters and place it at the bottom of the 7 ml scintillation vial. It is important to force the filter to the bottom of the vial so that the small volume of ethyl acetate (step 11) will completely cover the filter. 10. Allow the filter to dry before proceeding to the next step. Any ethanol remaining on the filter must be evaporated as it is a potent quencher in scintillation counting. 11. Add 0.5 ml of ethyl acetate to the scintillation vial. 12. Once the filter is dissolved, add 5 ml of scintillation cocktail, and vortex the vial briefly. 13. It is usually necessary to allow the sample to sit for as long as two days to maximize the dispersion of the radioactivity into the cocktail. Vortex briefly before scintillation counting. Rate of leucine incorporation (Leu incorp as nmol 1 ~h ') is calculated as follows: Leu incorp = {(dpm on sample filter)-(dpm in killed control)}/ incubation time/(2.22 × 10" d p m per tJCi) x (Leu specific activity as nmol per laCi) The factor 2.22 x 10" d p m per laCi converts the radioactivity (dpms) found on the filter to laCi, the unit of radioactivity used for the leucine specific activity. This specific activity is provided for each batch of leucine by the manufacturer and converts radioactivity (tJCi) to moles of leucine incorporated.
M E A S U R I N G 3H-LEUCINE I N C O R P O R A T I O N BYTHE MICROCENTRIFUGE METHOD Principle The following procedure, which was originally proposed by Smith and Azam (1992), is nearly the same as the flter method just described. The critical differences are that the incubation and radioassaying are both done in a 2.0 ml microcentrifuge tube. Also, the radioactivity incorporated into the microbial biomass is collected by centrifugation, not filtration. As a result, the amount of radioactivity and the volume of sample and scintillation cocktail are much smaller than used in the filter method. Also, the processing time and variability among replicates are often better with the microcentrifuge method. Finall),, the 'blank' or radioactivity in the killed controls is usually much lower for the microcentrifuge method (especially when TdR incorporation is measured), which allows much lower rates to be measured in shorter incubation times. This is especially important w h e n samples from deep waters with low activity are studied. 232
The m e t h o d described below has been extensively used in seawater and to a lesser extent in freshwaters. For freshwater systems, precipitation m a y be aided by adding humics (Kirschner and Velimirox; 1999) or NaC1 (to 3.5'7,, final concentration; D.C. Smith, personal communication), but other investigators do not add anything (J.J. Cole, personal communication). The need for additions to aid in precipitation should be assessed for each freshwater system u n d e r study. Additions are not needed for seawater studies.
Equipment and reagents • Tritiated leucine ([2,3,4-3H] leucine) as a stock solution with specific activity >60 nmol per Ci (I mCi ml ') (NEN, Amersham). • 2 ml microcentrifuge tubes. • Pipettes (e.g. Pipetman) that dispense volumes ranging from microliters (for the 3H-leucine) to milliliters. Repeating dispensers for the washes and scintillation cocktail. • Trichloroacetic acid (TCA) in a concentrated solution (100% w/v) and as a wash solution (5%). • Ethanol, 80%. • Aspirator, which can be constructed with tubing and a vacuum pump. • Scintillation cocktail (7 ml). Ultima-Gold (Packard Instruments) was found to be the optimal cocktail (H.W. Ducklow, personal communication). • Vortexer. • Microcentrifuge (e.g. Eppendorf). • 7 ml plastic scintillation vials (can be re-used) as carriers for the microcentrifuge tube. Scintillation vials from some manufacturers are smaller than others which makes it difficult to remove the microcentrifuge tube. • Scintillation counter (e.g. Beckman,Wallac). • Appropriate containers for radioactive corrosive liquids and radioactive solids.
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The same general comments about sterile technique, contamination, and gloves that apply to the filter m e t h o d also apply to the microcentrifuge method. 1. Add an appropriate v o l u m e of ['H] leucine to each microfuge tube before the sample is added. The tubes need not be washed before use. The stock [~H] leucine will need to be diluted such that a small but reasonable v o l u m e (between 2 and 5 Ill) of [:'HI leucine can be a d d e d to each tube. As with the filter method, the final concentration of [~H] leucine should be 20 nM, but this can be tested as described above. Killed controls are prepared by adding 89 pl of 100~ TCA to selected tubes. It is convenient to mark one side of the tube (cap and top edge) for positioning in the microcentrifuge (see step 7).
233
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2. Add sample water (1.7 ml) to each microfuge tube and shake the tube by inverting it. Duplicates or triplicates should be prepared per sample. 3. Incubate the samples at the i~l sihl temperature for an appropriate time period as described for the filter method. 4. After incubation, add 89 t~1of 100% TCA to each sample, except for the killed controls. (For measuring TdR incorporation, the samples must be cooled to 4°C by placing the tubes in ice before addition of the TCA.) 5. Place the tubes in the microcentrifuge with the mark on the outside, i.e. the side where the pellet will eventually form. Centrifuge the samples at the maximum speed of the microcentrifuge for 10 min. 6. After centrifugation, remove the supernatant by aspiration, being careful to avoid the pellet which is on the marked side. 7. Add l ml of ice-cold TCA to each tube, vortex, and repeat the centrifugation step. 8. Remove the TCA by aspiration as described in step 7 and add 1.0 ml of ice-cold 80% ethanol. Centrifuge again. 9. Remove the ethanol by aspiration. 10. Allow the pellet to dry completely because any remaining ethanol will cause quenching during liquid scintillation counting and lead to erratic results. 11. Add 1 ml of scintillation cocktail to the microcentrifuge tube and then vortex briefly. 12. Place the microcentrifuge tubes in plastic 7 ml scintillation vials and radioassay. 13. It is usually necessary to allow the sample to sit for as long as two days to maximize the dispersion of the radioactivity into the cocktail. Vortex briefly before scintillation counting. Rates of leucine incorporation (Leu incorp as nmol 1 ' h 1)are calculated using the same equation as used for the filter method.
CONCLUSIONS ideally both leucine and thymidine methods should be used because they provide independent estimates of bacterial production. The dual label method with I4C-leucine and ~H-thymidine allows both incorporation rates to be estimated in a single incubation tube (Chin-Leo and Kirchman, 1988). Some investigators have used leucine as a measure of biomass production (gC per liter per day) and thymidine for cell production (cells per liter per day). Their rationale is that leucine measures protein synthesis, i.e. biomass production, whereas thymidine reflects DNA synthesis, i.e. cell production. During balanced growth, however, the two rates have to be equal (when converted to equivalent units) because for sustained periods, cells cannot increase mass without division nor can 234
they divide without making new biomass. For this reason, leucine and thymidine incorporation rates cannot diverge for long time periods (greater than a couple of generation times or roughly a few days) or over large geographic areas. In practice, leucine and thymidine incorporation rates usually covary and are highly correlated. But since bacterial growth is not necessarily balanced, rates of leucine and thymidine incorporation may diverge and not be correlated. The difference between leucine and thymidine incorporation may be informative, if coupled with other measurements of the biogeochemical environment. Rates of leucine and thymidine incorporation could also diverge because the relationship between incorporation rates and bacterial production, which is reflected in conversion factors, may change. For example, changes in DNA content per cell, perhaps due to changes in bacterial species composition, would lead to variation in thymidine incorporation without changes in the actual rate of biomass production. Changes in isotope dilution is probably the biggest unknown in calculating production from leucine incorporation. The problem of picking the correct conversion factor is the difficult part of using either leucine or thymidine incorporation as a measure of bacterial production. Using 'empirical' conversion factors potentially is a solution, but it is far from perfect. Ducklow (2000) provides the most recent review of these questions. It should be emphasized that often rates of leucine and thymidine incorporation alone, e.g. moles of leucine incorporated per liter per day, are sufficient for addressing ecological questions. Similarly, often an estimate of incorporation rates per cell is an adequate index of bacterial growth rates. Even when using just incorporation rates, however, one conceptional problem with the leucine method should be mentioned. Unlike DNA, protein can 'turn over', i.e. protein is degraded within a cell and new protein is synthesized, in prokaryotes protein turnover is generally thought to be negligible, but conceivably it is substantial compared to total protein synthesis when cells are growing very slowly, i.e. the case with some aquatic environments. If protein turnover is substantial, then leucine incorporation would overestimate bacterial production. In the extreme case of high protein turnover and zero net protein synthesis (and thus zero biomass production), PHlleucine would be incorporated into the new protein but no radioactivity would be lost during the degradation of the old (non-radioactive) protein. Thus, the leucine method would indicate some positive rate of biomass production when in fact there was none. The single study of protein turnover in an aquatic environment (Kirchman et al., 1986) did not find substantial rates, but more work is needed on this question. The other main conceptual problem with both the leucine and thymidine method is that it measures total production of the entire community. It is now well recognized that the heterotrophic bacterial community is very diverse and that the role in biogeochemical cycles for each member of this community is likely to differ. For this reason, there is a need to develop methods for measuring biomass production and growth rates for specific members or groups within the bacterial community. Some progress has been made (Kemp et al., 1993; Urbach et al., 1999), but more 235
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w o r k is n e e d e d . It is conceivable that bacterial p r o d u c t i o n will be m o r e accurately estimated b y a d d i n g u p the c o n t r i b u t i o n s b y the m a j o r bacterial g r o u p s rather t h a n t r y i n g to m e a s u r e a c o m m u n i t y rate, as described here.
References Chin-Leo, G. and Kirchman, D. L. (1988). Estimating bacterial production in marine waters from the simultaneous incorporation of thymidine and leucine. Appl. Environ. Microbiol. 54, 1934-1939. del Giorgio, P. A., Prairie, Y. T. and Bird, D. E (1997). Coupling between rates of bacterial production and the abundance of metabolically active bacteria in lakes, enumerated using CTC reduction and flow cytometry. Microbial Ecol. 34, 144-154. Ducklow, H. W. (2000). Bacterial production and biomass in the oceans. In: Microbial Ecology o(tlle Oceans (D. L. Kirchman, Ed.). John Wiley and Sons, New York. Fuhrman, J. A. and Azam, F. (1980). Bacterioplankton secondary production estimates for coastal waters of British Columbia, Antarctica, and California. Appl. Environ. Microbiol. 39, 1085-1095. Euhrman, J. A. and Azam, E (1982). Thymidine incorporation as a measure of heterotrophic bacterioplankton production in marine surface waters: evaluation and field results. Mar. Biol. 66, 109-120. Fukuda, R., Ogawa, H., Nagata, T. and Koike, 1. (1998). Direct determination of carbon and nitrogen contents of natural bacterial assemblages in marine environments. Appl. Environ. Microbiol. 64, 3352-3358. Kemp, P. E, Lee, S. and LaRoche, J. (1993). Estimating the growth rate of slowly growing marine bacteria from RNA content. Appl. Envi~vn. Microbiol. 59, 2594-2601. Kirchman, D. L. (1993). Leucine incorporation as a measure of biomass production by heterotrophic bacteria. In: Current Methods in Aquatic Microbial Ecology (P. Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds). Lewis Publishers, Boca Raton, EL. Kirchman, D. L., K'nees, E. and Hodson, R. E. (1985). Leucine incorporation and its potential as a measure of protein synthesis by bacteria in natural aquatic systems. Appl. Environ. Microbiol. 49, 599-607. Kirchman, D. L., Newell, S. Y. and Hodson, R. E. (1986). Incorporation versus biosynthesis of leucine: implications for measuring rates of protein synthesis and biomass production by bacteria in marine systems. Mar. Ecol. Prog. Set. 32, 47-59. Kirschner, A. K. T. and Velimirov, B. (1999). Modification of the H-3-1eucine centrifugation method for determining bacterial protein synthesis in freshwater samples. Aquatic Microbial Ecol. 17, 201-206. Sherr, B. E, del Giorgio, P. and Sherr, E. B. (1999). Estimating abundance and single-cell characteristics of respiring bacteria via the redox dye CTC. Aquatic Microbial Ecol. 18, 117-131. Simon, M. and Azam, E (1989). Protein content and protein synthesis rates of planktonic marine bacteria. Mar. Ecol. Prog. Set. 51, 201-213. Smith, D. C. and Azam, E (1992). A sirnple, economical method for measuring bacterial protein synthesis in seawater using ~H-leucine. Mar. Micn)b. Food Webs 6, 107-114. Urbach, E., Vergin, K. L. and Giovannoni, S. J. (1999). hnmunochemical detection and isolation of DNA from metabolically active bacteria. Appl. Environ. Microbiol. 65, 1207-1213.
236
List of suppliers The following is a selection of companies. For most products, alternative suppliers are available.
Amersham Life Science, Inc. 26111 Miles Road Cleveland, OH 44128, USA Teh 1-216-765-5000 Fax: 1-216-464-5075 http://www.amersham.co.uk Radiochemicals ([~H]Leu, [~H]TdR).
Packard Instrument Company 800 Research Parkway Meriden, CT 06450, USA Teh 1-800-323-1891 USA Only 1-203-238-2351 Fax: 1-203-639-2172 E-maih
[email protected] Scintillation counter; scintillation cocktail.
Beckman Instruments, Inc. 2500 Harbor Blvd. Fullerton, CA 92634-3100, USA Teh 1-714-871-4848 Fax: 1-714-773-8283 1-800-643-4366
Rainin Instrument Co., Inc. Mack Road, Box 4026 Woburn, MA 01888-4026, USA Tel: 617-935-3050 Pipetters.
Scintillation counter.
Millipore Corporation 80 Ashby Road Bedford, MA 01730, USA Teh 1-800- 645-5476 1-617-275-9200 Fax: (508) 624-8873
Wallac Inc., Berthold 9238 Gaither Road Gaithersburg, MD 20877, USA Tel: 1-301-963-3200 1-800-638-6692 Fax: 1-301-963-7780 e-mail:
[email protected]
Filters.
Scintillation counter.
NEN Life Science Products 549 Albany Street Boston, MA 02118, USA Tel: 1-800-551-2121; 1-617-482-9595 Fax: 1-617-482-1380 Radiochemicals ([~H]Leu, [~H]TdR).
237
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13 Phosphorus Cycle in Seawater: Dissolved and Particulate Pool Inventories and Selected Phosphorus Fluxes DM Karl and KM Bj6rkman Department of Oceanography,School of Ocean and Earth Scienceand Technology,University of Hawaii, Honolulu, HI 96822, USA
CONTENTS General introduction Detection of phosphorus and P-containing compounds in seawater High-sensitivity, high-specificity assay for Pi and measurements of TDP Particulate phosphorus Pi uptake/regeneration and DOP production/utilization rates Intracellular ATP pool turnover and biologically available P
######
GENERAL INTRODUCTION Phosphorus (P) is an essential macronutrient for all living organisms; life is truly built around P (deDuve, 1991). In the sea, P exists in both dissolved and particulate pools with inorganic as well as organic origins. The uptake, remineralization/hydrolysis and exchanges (by both physical and biological processes) of these various pools are the essential components of the marine P cycle (Figure 13.1). Compared to the much more extensive investigations of carbon (C) and nitrogen (N) dynamics in the sea, P pool inventories and fluxes are less well documented although no less important. Herein, we present the principle and stepwise procedures for the accurate estimation of (1) dissolved orthophosphate (HPO~ 2 ; hereafter referred to as Pi), soluble reactive P (SRP), total dissolved P (TDP) and particulate P (PP) as minimal constraints on P pool inventories, (2) dissolved and particulate adenosine-5'-triphosphate concentrations (D-ATP and P-ATP, respectively), (3) Pi uptake/regeneration and dissolved organic P (DOP) production rates using radiolabeled Pi (~2p or s~p) precursors and (4) turnover rate estimation of the c~-P and y-P of intracellular ATP. The latter
METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0-12-521530 4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any form reserved
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Atm deposition (wet and dry)
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Diffusion, mixing, active transport
export
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hydrolysiS" Diffusion, mixing, active transport Particplate
I Low density Exportflux upward flux (gravitational and active migrations)
Figure 13.1. Schematic representation of the open ocean P-cycle showing the various sources and sinks of inorganic and organic P pools, including biotic and abiotic interconversions. The large rectangle in the center represents the upper water column TDP pool comprised of Pi, inorganic polyphosphate and a broad spectrum of largely uncharacterized DOP. Ectoenzymatic activity (Ecto) is critical for microbial assimilation of selected TDP compounds. Particulate P, which includes all viable microorganisms, sustains the P-cycle by assimilating and regenerating Pi, producing and hydrolyzing selected non-P/ P, especially DOP compounds, and by supporting net particulate matter production and export. Atmospheric deposition, horizontal transport and the upward flux of low density organic P compounds are generally poorly constrained processes in most marine habitats. Phosphine (PHi), shown at the right, is the most reduced form of P in the biosphere and is generally negligible except under very unusual, highly reduced conditions.
240
measurements provide information on microbial growth rate, total energy flux and the biologically available P (BAP) pool. While this by no means represents a comprehensive investigation of P cycle processes (see Table 13.1), these methods collectively provide a protocol suite that is suitable for an initial study of P dynamics in selected marine habitats.
D E T E C T I O N OF P H O S P H O R U S A N D P - C O N T A I N I N G C O M P O U N D S IN S E A W A T E R The analysis of dissolved and particulate P-containing compounds in seawater is neither simple nor straightforward. Strickland and Parsons (1972) defined eight different operational classes of P compounds based on reactivity with the acidic molybdate reagents, ease of hydrolysis and particle size. These range from 'inorganic, soluble and reactive,' presumably Pi, through 'enzyme hydrolyzable phosphate' (Pi released following treatment with the enzyme alkaline phosphomonoesterase), to 'inorganic, particulate and unreactive' (presumably P-containing minerals). Some of the operationally-defined pools have no convenient analytical method of determination while others can be estimated only as the difference between two operational classes with partially overlapping specificity. Only a very few specific compounds or compound classes can be readily detected at the low concentrations typically found in seawater (Table 13.1). There is likely to be a broad spectrum of P-containing compounds in seawater. Our inability to completely characterize these various dissolved and particulate pools currently limits further progress towards a comprehensive understanding of the marine P cycle. Even the most routine analytical method employed for Pi concentration measurement in seawater fails to provide an accurate estimate, especially for habitats where the DOP pool exceeds the dissolved inorganic P pool (e.g. most subtropical and tropical surface waters). Quantitative analyses of P in seawater have traditionally relied upon the formation of a 12-molybdophosphoric acid (12-MPA) complex and its subsequent reduction to yield a highly colored blue solution, the extinction of which is measured by absorption spectrophotometry (Fiske and Subbarow, 1925; Murphy and Riley, 1962). Over the years, numerous improvements have been introduced to the basic method so that substantial variability now exists in the conditions used for color development, final reduction of the 12-MPA complex, and the treatment of potentially interfering compounds. Although the stepwise chemical procedures for P determination are straightforward and fully amenable to automated analysis, there are many complexities, both analytical and conceptual, inherent in measuring and interpreting P concentrations in seawater (Tarapchak, 1983). For example, the soluble reactive P (SRP) pool measured by the standard Murphy-Riley procedure is not necessarily equivalent to the concentration of Pi, but may also include DOP compounds that are hydrolyzed 241
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under the acidic reaction conditions. Significant differences between SRP and Pi have been observed for nearly every natural aquatic ecosystem where more rigorous and specific methods of Pi analysis have been employed (Kuenzler and Ketchum, 1962; Jones and Spencer, 1963; Rigler, 1968; Pettersson, 1979; Karl and Tien, 1997; Thomson-Bulldis and Karl, 1998), and even different SRP methods return unequal estimates of 'Pi' when employed with common seawater samples (Karl and Tien, 1997). As discussed below, accurate determination of Pi is absolutely essential for the application of ~Pi or ~;Pi tracer studies if mass flux estimation (Pi uptake or Pi regeneration rates) is the experimental objective. Most investigators measure both SRP and TDP; the latter is also quantified by the molybdenum blue reaction following hydrolysis (Table 13.1). Only the nitrate oxidation method provides a quantitative recovery of total P, including polyphosphates and phosphonates (Cembella et al., 1986). Monaghan and Ruttenberg (1999) have recently re-evaluated the efficacy of several methods for TDP estimation in coastal waters, and recommend the Sol6rzano and Sharp (1980) procedure. It is conceivable, even likely, that the behavior of these various methods is site-specific and that the efficiency of soluble non-reactive P (SNP) hydrolysis to SRP is dependent upon the chemical composition of the P pool in a given habitat.
e,e,e,e,e,~, H I G H - S E N S I T I V I T Y , H I G H - S P E C I F I C I T Y A S S A Y FOR Pi A N D M E A S U R E M E N T S OF T D P Principle In most oligotrophic oceanic environments, Pi determinations are unreliable because the ambient concentrations approach the limit of analytical detection (-20-40 nM) for the standard Murphy-Riley method. At these low concentrations, the variability of replicate determinations can be +25%, or greater. Consequently, several modifications and new analytical procedures have been introduced to improve the detection limit and precision of environmental Pi analyses (Table 13.1). The magnesium-induced coprecipitation (MAGIC) method for the precise determination of Pi, described by Karl and colleagues (Karl and Tien, 1992; Thomson-Bulldis and Karl, 1998) can be used to eliminate interference from non-Pi, P-containing compounds thereby providing a more accurate estimate of Pi that is necessary for reliable P-cycle flux determinations (Table 13.1). The method relies on the quantitative removal of P from solution by in vitro formation of brucite [Mg(OH)2], initiated by the addition of NaOH. The precipitate is collected by centrifugation and dissolved in HC1 for Pi determination by the standard Murphy-Riley molybdenum blue reaction. This MAGIC procedure can be used to routinely effect a Pi pre-concentration ranging from 2-100-fold, thereby providing a method which can reliably detect subnanomolar concentrations in seawater. The method is highly reproducible; typical estimates of precision for triplicate SRP determinations in the 10-100 nM 247
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range are from 1 to 3%. The method is versatile and has m a n y potential ecological applications.
Pi Determination by the modified MAGIC technique Equipment and reagents • • • • • • • • •
• • •
Acid cleaned high-density polyethylene (HDPE) sampling bottles Polypropylene centrifuge tubes, 50 ml (Coming) Plastic 50 ml pipettes for sample dispensing Automatic pipettes or repipetter for reagent dispensing Centrifuge with capacity to hold 50 ml tubes Spectrophotometer, Beckman DU 640, or equivalent, and cuvette cells (I0 cm) Sodium hydroxide I H (NaOH, Fluka Biochemica #71689) Hydrochloric acid 0. I H (Fisher trace metal grade HCI #A508-212) Arsenate reducing mixture: I. Sulfuric acid 3.5 N (H2504 Fisher #A300-212) 2. Sodium metabisulfite 14% wlv (Na2OsS2, Fluka Biochemica #71930) 3. Sodium thiosulfate 1.4% wlv (Na203S2, Fluka Biochemica #72_049) All solutions are made up in distilled, deionized water (DDW). Sodium metabisulfite must be freshly prepared. Mix reagents I and 2, add 3 in the volumetric proportions 2:4:4. Primary standard potassium phosphate (KH2PO4); prepare dilution series 0.015-0.5 IJM, including blank Certified reference material (CSK-nutrient element, Wako Chemicals #27874099) Molybdenum blue reaction mixture: I. Sulfuric acid 5 N (H2SO4, Fisher #A300-212_) 2. Ammonium molybdate 30% wlv ((NH4)6MoTO24 • 4H20, Fisher #A674-S00) 3. Ascorbic acid S.4% wlv (Fisher #A6 I- 100) 4. Potassium antimonyl tartrate 0.136% wlv (KSbC4H407 • 0.5 1-120, Mallinckrodt #2388) All solutions are made up in DDW. Ascorbic acid must be freshly prepared, or store frozen prior to use. All other solutions are stable for months, or longer. Mix reagents I and 2, add 3 and 4 in the volumetric proportions 5:2:2:1.
Assay 1. 2.
The stored, frozen samples are thawed in a water bath to room temperature. Triplicate 50ml sample aliquots are placed into new 50ml polypropylene centrifuge tubes using a 50 ml plastic pipette. Work from anticipated low to high concentrations (i.e. from surface to deep) or thoroughly rinse pipette in distilled water between samples.
248
3. 4. 5. 6.
7.
8. 9.
Add 150 pl 1 M NaOH to each 50 ml aliquot (0.3%, v/v). Cap tubes tightly, and invert to mix the floc produced by the NaOH addition. Centrifuge at 1000g at room temperature for 60 rain. Aspirate the supernatant using a clean Pasteur pipette attached to a vacuum line and collection flask. Take care not to disturb the small pellet. For a routine five-fold sample concentration, add 10 ml of 0.01 M HC1 to each tube and vortex to dissolve the pellet. Greater concentration factors up to and exceeding 100-fold can be effected by decreasing the final volume of HC1, by increasing the initial seawater volume, by increasing the acidity of the HC1, or all three. Add 1 ml reduction mix to each 10 ml sample. Vortex and allow to react for 15+1 min. After 15+1 min add 1 ml molybdenum blue reaction mix to each 10 ml sample and vortex to mix. Wait at least 15rain at room temperature to ensure full color development.
Note: Standards are prepared in surface seawater and treated the same as the samples. Blank samples are prepared from phosphate-free seawater (i.e. supernatant from MAGIC treated samples). Samples and standards are read at 880 nm in a 10 cm cell.
Optionah Tracer addition of ~Pi (or ~'Pi) added to the original sample prior to NaOH addition can be used to precisely determine the recovery of Pi for each subsample. While this is not necessary for routine use, it does provide the most accurate and precise estimation of Pi.
The measurement of TDP is also operationally-defined; typically a high-intensity UV photooxidation (Armstrong et al., 1966) or high temperature wet persulfate oxidation (Menzel and Corwin, 1965) or a combined procedure (Ridal and Moore, 1990) pre-treatment is used to convert SNP to Pi for subsequent analysis by the standard molybdenum blue assay. However, it is well known that certain P-containing compounds (e.g. inorganic polyphosphates, nucleotide di- and triphosphates) are not quantitatively recovered by standard UV photooxidation procedures; neither method quantitatively recovers P from phosphonate compounds. Depending upon the methods used, the difference between the measurement of TDP and either Pi or SRP will be termed SNP (i.e. SNP = [TDP][SRP]) or non-P/ P (N-P/ P = [TDPI-[Pi]), where SNP ~ non-Pi P (Thomson-Bulldis and Karl, 1998). Furthermore, the MAGIC technique provides the opportunity for a direct rather than an indirect measurement of SNP (technically, non-Pi P) by an initial quantitative removal of Pi, followed by sample hydrolysis-oxidation and Pi determination (Thomson-Bulldis and Karl, 1998). This is especially useful for measure249
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ments of non-P/P in waters where Pi is the dominant form, for example in the deep sea. As emphasized previously, there is no a priori relationship between these operationally-defined pools and the more ecologically-relevant BAP pool.
TDP determination by persulfate oxidation and MAGIC Equipment and reagents • Acid-cleaned high-density polyethylene (HDPE) sampling bottles • Acid-cleaned autoclavable glass centrifuge tubes 50 ml with Teflon lined caps (Kimble) • Plastic 50 ml pipettes for sample dispensing • Automatic pipettes or repipetter for reagent dispensing • Centrifuge with capacity to hold 50 ml tubes • Autoclave • Sodium hydroxide I M (Na©H, Fluka Biochemica #71689) • Hydrochloric acid 0. I M (Fisher trace metal grade HCI #A508-212) • Potassium persulfate 5% w/v (K2S208,Fisher #P281) • Primary standard potassium phosphate (KH2PO,); prepare dilution series 0.015-0.5 lUM,including blank • Spectrophotometer Beckman DU 640, or equivalent, and cuvette cells (10 cm) • Certified reference material, arsenate reducing mixture and molybdenum blue reagents (all as above)
Assay 1. The stored, frozen samples are thawed in a water bath to room temperature. Typically, subsamples for TDP analysis are processed along with Pi determinations (see above). 2. Triplicate 40 ml sample aliquots are placed into 50 ml acid-washed glass centrifuge tubes using a 50 ml plastic pipette. Work from anticipated low to high concentrations (i.e. from surface to deep). 3. Add 6.4 ml potassium persulfate solution to each tube. Cap tightly. 4. Autoclave 121°C, 30 min. 5. Allow samples to cool to room temperature. 6. Add 3.3 ml I M NaOH to each sample. 7. Cap tubes tightly, and invert to mix the floc produced by the NaOH addition. 8. Centrifuge at 1000g at room temperature for 60 min. 9. Aspirate the supernatant using a clean Pasteur pipette attached to a vacuum line and collection flask. Take care not to disturb the pellet. 10. Add 9.0 ml of 0.1 M HC1 to each tube and vortex to dissolve the pellet completely.
250
11. A d d 1 ml reduction mix to each 10 ml sample. Vortex and allow to react for 15+1 min. 12. After 15+1 min add 1 ml m o l y b d e n u m blue reaction mix to each 10 ml sample and vortex to mix. Allow to react for at least 15 min at room temperature to ensure full color development. Samples and standards are read at 880 n m in 10 cm cells. N~te: Standards are prepared in surface seawater and treated the same as samples. Blank samples are prepared from artificial seawater (Instant Ocean) filtered through 0.21Jm m e m b r a n e filter. After a MAGIC treatment (2.5% v / v 1 M N a O H addition, centrifuge at 1000g for 1 h), 40 ml aliquots of the supernatant are transferred to glass tubes and treated as regular samples from step 3, above.
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PARTICULATE PHOSPHORUS
Rationale We define 'particles,' operationally, as those materials that are retained by a W h a t m a n micro-fine glass fiber filter, catalog # G F / F 0.7 ~m nominal retention (Sheldon, 1972), acknowledging the fact that some particulate material is k n o w n to pass through this filter. The particle retention characteristics are also k n o w n to vary with filter loading and, hence, m a y not be constant even during the filtration of a h o m o g e n e o u s water sample. In presenting our methods, we have focused on total elemental mass and applications to mass fluxes without consideration as to the chemical form of the PP. Consequently, we derive estimates that cannot be assumed to be equivalent to the organically-bound fraction as they m a y include inorganic PP as well. More specialized methods do exist for the measuremerits of lipid-P and nucleic acid-P in the isolated particulate matter (Table 13.l), but these will not be discussed. The procedure presented herein is a modification of one used by the Hawaii Ocean Time-series (HOT) p r o g r a m at the University of Hawaii and relies on the release of organically-bound p h o s p h o r u s c o m p o u n d s as Pi, by high-temperature combustion at 450°C. The Pi is then extracted with HC1 at 60°C and neutralized with NaOH. The liberated Pi is then measured using the M u r p h y - R i l e y procedure described above (see Pi determination). This procedure measures all forms of reactive p h o s p h o r u s released by combustion and acid hydrolysis. Application of this m e t h o d for PP inventory estimation and for PP fluxes using particle interceptor traps has been presented elsewhere (Karl et al., 1991, 1996).
251
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Particulate phosphorus determination Equipment and reagents • • • • • • • • • • • •
Acid-cleaned polyethylene (PE) bottles (I 2 I) Low pressure (4-7 psi nitrogen gas) or vacuum filtration apparatus Filter holder assembly for in-line pressure filtration Muffle furnace (450°C) Drying oven (60°C) Combusted (450°C, 4.5 h), HCI (I M) washed, glass tubes 12 x 75 mm Combusted, HCI (I M) rinsed GF/F filters, 25 mm Combusted aluminum foil Clean forceps Automatic pipettes or repipetter for reagent dispensing Hydrochloric acid 0.15 N Certified reference material, phosphate standard solutions, molybdenum blue reaction mixture and equipment (all as above for Pi determination)
Assay 1.
2.
3.
4. 5. 6. 7. 8. 9.
Seawater is collected using appropriate sampling techniques. If necessary, samples are pre-screened through a 202 pm Nitex screen to remove large zooplankton. Samples are filtered through a combusted, acid-rinsed GF/F filter. This can be done by pressure filtration using in-line filter holders and applying 4-7 psi gas pressure (N2) or by vacuum filtration. Following filtration the filter is transferred into acid-cleaned, combusted glass tubes using clean forceps. Combusted, acidwashed GF/F filters serve as blank samples. The glass tubes are covered with a piece of combusted foil. The foil is secured with label tape and stored frozen (-20°C). Combust samples in muffle furnace (450°C, 4.5 h; remove tape before combusting). After cooling to room temperature, add 10 ml 0.15 N HC1; heat to 60°C for I h. Adjust volume to 10 ml, and centrifuge at 1000g for 15 min. Transfer 5 ml of supernatant into acid-washed, combusted glass tube. Add 0.5 ml of the molybdenum blue reaction mixture to each sample; mix and allow to react for i h.
Note: Standards are prepared in 0.15 N HC1 and treated the same as samples. Samples and standards are read at 880 nm in 1 cm cells. Filter blanks are also included.
252
Measurements of particulate and dissolved ATP Rationale: P a r t i c u l a t e A T P as a microbial biomass indicator
The phylogenic diversity of microorganisms in the sea and the generally high percentage of non-living to living C, N and P complicates a straightforward assessment of total microbial biomass despite the fundamental importance of this parameter for many ecological studies. Measurements of PP (see above) can be used to provide an upper bound on biomass-P, but in most open ocean surface waters and in all subeuphotic zone waters of the global ocean non-living components dominate the standing stock of POM. It would, therefore, be desirable to have a reliable biochemical or molecular biomarker for life, one that is: (1) present in all living cells and readily metabolized, hydrolyzed or otherwise decomposed following cell death, (2) present in a fixed, constant percentage relative to total cell mass regardless of environmental or physiological conditions; and (3) easily extracted and purified (if necessary) from seawater and conveniently measured. Levin et al. (1964) first suggested that ATP, the common energy currency of all living cells, might be a sensitive indicator for the presence of living microorganisms in natural samples. Two years later, HolmHansen and Booth (1966) proposed that ATP measurements could be used to estimate total microbial biomass in marine ecosystems. These pioneering efforts evolved into a simple and extremely sensitive assay that has widescale use in marine microbiology. Without exception, all living organisms contain ATP, so the basis for the use of this molecular biomarker is well-founded (Karl and Dobbs, 1998). The measurement of P-ATP is described in detail elsewhere (Karl, 1993).
Rationale: Dissolved A T P as a ' m o d e l ' D O P c o m p o u n d
Significant concentrations of D-ATP have been detected in marine (Azam and Hodson, 1977; Nawrocki and Karl, 1989) and freshwater (Riemann, 1979; Maki et al., 1983) environments. It is unknown whether this D-ATP interferes with the conventional measurement of P-ATP in aquatic environments, but since most water samples are generally concentrated onto filters before extraction, the relative contribution from D-ATP should be minimal. The much more interesting questions with regard to tile ubiquity of D-ATP in natural aquatic ecosystems are: Why? And by what mechanisms? Given the fundamental role of ATP in cellular energetics, it is unlikely that D-ATP is derived directly from healthy cells by excretion or exudation. D-ATP production is probably a manifestation of microbial cell death by grazing or autolysis. D-ATP is also a readily bioavailable substrate; marine microorganisms can and do assimilate D-ATP, in part, for P and, in part, for purine salvage and nucleic acid synthesis. Consequently, D-ATP concentration measurements coupled with D-AT~-~P/~P (or ~H/14C radiolabeled substrates) can be used to estimate D-ATP pool turnover and mass flux through marine microbial assemblages (Azam and Hodson, 1977). 253
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D-ATP is probably the only organic-P compound that can be measured at ambient pool concentrations and can be tracked through the microbial food web maze. In this regard, and given its fundamental role in cellular bioenergetics and biosynthesis, ATP might be considered a model compound in studies of the marine P cycle. In addition to D-ATP, dissolved ADP and AMP (McGrath and Sullivan, 1981), dissolved cyclic-3', 5'-AMP (c-AMP; Ammerman and Azam, 1981) and dissolved guanosine-5'-triphosphate (Bj6rkman and Karl, 2000) have also been detected in seawater. At the present time, it is not known whether these related intracellular compounds have similar sources and sinks in the marine environment.
Dissolved ATP Equipment and reagents • • • • • • •
Acid-cleaned high-density polyethylene (HDPE) sampling bottles Vacuum filtration manifold Membrane filters (0.2 IJm, Poretics, Nuclepore) Adjustable automatic pipettes (10-100 lUl, 100-1000 IJl, I-5 ml) Sodium hydroxide (I M NaOH, Fluka Biochemika #71689) Hydrochloric acid (5 M HCI, Fisher #AI44S-212) Modified firefly lantern extract (FLE) mixture for high-sensitivity bioluminescence assay: I. Firefly lantern extract, freeze dried (Sigma Chemical Co,# FLE-50) 2. -Iris 20 mM, pH 7.4 (Sigma Chemical Co,#T4003) 3. Arsenate buffer 0.1 M, pH 7.4 (Na2HAsO4, Mallinckrodt #7384) 4. Dithiothreitol 0.1 M (DTT; Sigma Chemical Co,#D-9779) 5. Luciferin 3.3 mM (Sigma Chemical Co,#L-6882) The FLE is reconstituted in 5 ml D D W per 50 mg vial and allowed to age at room temperature for 6-12 h prior to diluting with equal volumes of solutions 2 and 3 to a final volume 75 ml. The FLE mixture is filtered through a GF/F filter to remove solids. Add I% v/v DTT and luciferin, respectively, to the FLE mixture. • ATP photometer or equivalent light detection instrument, preferably equipped with an automatic injection unit and interfaced with a computer for calculations of peak height and the integrated counts of the light emission decay curve.
Assay 1.
Seawater samples are filtered through 0.2pro polycarbonate membrane filters and 450 ml of the filtrate transferred to HDPE bottles and stored frozen (-20°C) until further processed. The volume used is determined by the expected dissolved ATP concentration; smaller sample volumes can successfully be used with waters from eutrophic regions.
254
2. 3. 4. 5.
6.
7.
Thaw samples to room temperature in water bath. Transfer to clean transparent bottles. Add 2.25ml 1M NaOH (0.5% v/v) to 450ml sample. Mix thoroughly. Centrifuge at 1000g for 60 rain at room temperature. Aspirate supernatant and discard. (If a centrifuge that can accommodate large volume samples is not available the floc can be settled passively for 3 h. Aspirate the overlying volume until approximately 50 ml remains. Transfer this suspended floc to clean 50 ml polypropylene centrifuge tubes and centrifuge as in step #4.) Dissolve pellet by adding 5 M HC1 until solution just turns from milky to clear (pH 7-8), usually 100-200 pl. Adjust volume with Tris buffer (20 mM, pH 7.4) to 2 ml. Place 100 pl sample in reaction vial and insert in photometer. Add 800 pl of the FLE reaction mixture and monitor the light emission.
Note: Standards are prepared in deep-sea water with low D-ATP content from stock solutions and treated the same as unknown samples. The blank value is deducted and ATP concentrations are determined from the slope of the standard curve using peak height bioluminescence counts.
Pi U P T A K E / R E G E N E R A T I O N PRODUCTION/UTILIZATION
AND DOP RATES
Rationale The use of stable and radioisotopic tracers to monitor and quantify the rates of microbial growth, metabolism and biogeochemical cycling of key elements and compounds has revolutionized the field of microbiological oceanography. In general, there are two major categories of isotope-based research: (1) the use of naturally occurring stable or radioactive isotopes in selected inorganic substrate pools or organic matter and (2) the use of exogenously supplied stable or radioactive isotopes. The former application, which includes the detection of the cosmogenic radiotracers 32p and ~P (Lal and Lee, 1988; Lal et al., 1988) are most useful for long-term (day to week) whole ecosystem studies. Two recent applications in the Sargasso Sea and Gulf of Maine have demonstrated the efficacy of using natural cosmogenic ~P/~;P radioisotopes in studies of the marine P cycle (Waser et al., 1996; Benitez-Nelson and Buesseler, 1999). The latter application, also including the use of exogenously supplied ~:P and ~;P-labeled compounds, is best suited for short-term (hours to day) studies of metabolic pathways, nutrient fluxes and organic tissue labeling patterns. The use of ;~P (or ~P) as a tracer for Pi uptake, incorporation and regeneration has been used extensively in oceanography to measure the growth and metabolic activities of algal and bacterial assemblages (Rigler, 1956; Watt and Hayes, 1963; Sorokin and Vyshkvartsev, 1974; Taft et al., 1975; 255
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Harrison et al., 1977; Perry and Eppley, 1981; Sorokin, 1985; Atkinson, 1987; see Table 13.1). Certain P radiotracer field experiments have employed either size fractionation treatments (Harrison et al., 1977), metabolic inhibitors (Krempin et al., 1981 ) or multiple labeled substrates (Cuhel et al., 1983) to separate algal and heterotrophic bacterial activities. Typically, the radiotracer is added as carrier-free :~-Pi (or ~Pi) in order to minimize perturbations that may be caused by Pi pool enrichment. If accurate determination of ~'Pi concentration, or preferably BA~'P, is obtained then ~:Pi ('~Pi) uptake rates can be extrapolated to mass fluxes. The key to providing a reliable mass flux is the accurate estimation of the BAP pool. If standard SRP measurements are used, P uptake rates and, hence, mass fluxes could be grossly overestimated or underestimated. The two radioactive isotopes of P, ~-~Pand -~P, have very different characteristics including half-life and radioactive decay properties. While both isotopes produce 13 particles, the energy of ~2pdecay is much greater than ~'P and the half-life is shorter (~P, E......= 1.710 MeV, half-life = 14.3 days; ~P, E...... = 0.249 MeV, half-life = 25.3 days). A consequence of the ,2p high energy 13 emission is a phenomenon known as Cerenkov radiation. This property provides a unique opportunity to detect and quantify the activity of ~'P in dissolved and particulate matter with reasonable counting efficiencies (~40%) in the absence of a scintillation fluor and, therefore, without chemical quench. This non-destructive counting method means that the entire sample can be recovered for subsequent analysis. In addition to Pi uptake and rates of P incorporation into PP or specific subcellular pools (e.g. lipids, nucleic acids; see Table 13.1), exogenous radiotracer experiments can also provide information on Pi regeneration rates (Harrison, 1983; Smith et al., 1985) and DOP production rates (Johannes, 1964; Lemasson and Pages, 1981; Orrett and Karl, 1987; Bj6rkman et al., 2000). As for Pi uptake, accurate Pi regeneration or DOP production estimation requires information on the ambient pool of BAP in the habitat under investigation. When combined with independent methods of DOP pool characterization, these radiotracer experiments can also be used to quantify the fluxes of specific DOP compounds or compound classes (e.g. nucleotides). A 'pulse-chase' experimental design (i.e. pre-incubation with ~ePior ~Pi radiotracer followed by the addition of a ten-fold excess of ~'Pi) could be employed to follow both net DOP production, during the labeling phase, and DO~P turnover in the postchase treatments. Finally, the addition of exogenous, :~P (or :~P)-radiolabeled DOP compounds (e.g. glucose-P, ATP, glycerol-P) can provide information on the turnover time of individual pools and potentially (if combined with direct measurements of ambient pool concentrations) on mass fluxes (production and utilization rates) through selected organic-P pools. Our present inability to measure any but a very few DOP compounds (a notable exception is ATP, as discussed above) limits the application of this experimental design.
256
Pi uptake and DOP production Equipment and reagents • Acid-cleaned high density polyethylene (HDPE) sampling bottles • Polypropylene centrifuge tubes 50 ml (Corning) • Polycarbonate incubation bottles • On-deck temperature- and light-controlled incubator, or in situ incubation array • Vacuum filtration manifold, filtration funnels and bases, vacuum pump • Membrane filters (0.2 IJm Poretics, Nuclepore) • Glass scintillation vials • Acid cleaned autoclavable glass centrifuge tubes 50 ml with teflon lined caps (Kimble) • Plastic pipettes (50 ml) for sample dispensing • Automatic pipettes or repipetter for reagent dispensing and sampling • Centrifuge with capacity to hold 50 ml tubes • Steam or electric autoclave • Radiolabeled orthophosphate (32Pi/33Pi) as a stock solution of approximately 18.5 MBq ml' (500 luCi ml ~') (ICN Radiochemicals) • Sodium hydroxide I M (NaOH, Fluka Biochemica #71689) • Potassium persulfate 5% w/v (K2S208,Fisher #P281) • Hydrochloric acid 0. I M (Fisher trace metal grade HCI, #A508-212) • Spectrophotometer Beckman DU 640, or equivalent, and cuvette cells (10 cm) • Liquid scintillation counter (LSC) • Arsenate reduction mixture, Pi standards, molybdenum blue reaction mixture and equipment (as above for Pi determination) Assay 1. Incubate freshly collected seawater samples tinder defined temperature and light conditions in the presence of the radiolabel. The radioactivity required depends on sample size and ambient BAP concentrations. 2. At predetermined incubation periods, subsamples are removed for total (1 ml whole water) and particulate radioactivity by filtration onto membrane filters. The whole water and filter samples are placed in scintillation vials and the filtrate collected and stored frozen (-20°C) in HDPE bottles. 3. The filters and whole water samples are counted by LSC. 4. The filtrate samples are thawed in a water bath to room temperature. 5. Triplicate 50 ml sample aliquots are placed into new 50 ml polypropylene centrifuge tubes using a 50 ml plastic pipette; 1 ml from each sample is placed directly into a scintillation vial for LSC counting of total ~P (~'P) activity. 6. Add 150 lal (0.3% v/v) 1 M NaOH to each 50 ml aliquot.
257
¢. m
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7. Cap tubes tightly, and invert to mix the floc produced by the NaOH addition. 8. Centrifuge at 1000g at room temperature for 60 min. 9. Remove 1 ml of supernatant for LSC counting of remaining radioactivity (DO~2P/DO~P). 10. Transfer 3 x 40 ml of the supernatant to 50 ml acid washed glass centrifuge tubes using a 50 ml plastic pipette. 11. Aspirate the remaining supernatant in the polypropylene tubes taking care not to disturb the small pellet (go to step # 20). 12. To each glass tube add 6.4 ml potassium persulfate solution. Cap tightly. 13. Autoclave 121°C, 30 min. 14. Allow samples to cool to room temperature. 15. Add 3.3 ml 1 M NaOH to each sample. 16. Cap tubes tightly, and invert to mix the floc produced by the NaOH addition. 17. Centrifuge at 1000g at room temperature for 60 rain. 18. Aspirate the supernatant using a clean Pasteur pipette attached to a vacuum line and a collection flask. 19. Add 9.0 ml of 0.1 M HC1 is added to each tube and shake to dissolve the pellet. 20. Add 1 ml reduction mix to each 10 ml sample. Measure Pi, as above. Note: From time course incubations the uptake rate of '~Pi, and the
turnover time of the Pi equivalent pool can be calculated. If the Pi (SRP or BAP pool) is known the mass flux of P can be calculated from the turnover time and the known concentration of the relevant P-pool. The production of DOP is estimated from the increase in ~-'Pactivity in the supernatant over time and by assuming that the DOP formed has the same specific activity as the initial Pi. The production rate of DOP can be calculated and the average turnover time for the DOP pool assessed with the knowledge of the DOP pool size derived from the DOP concentrations in the samples.
e4l,~,4,ee I N T R A C E L L U L A R A T P P O O L T U R N O V E R B I O L O G I C A L L Y A V A I LABLE P
AND
Principle: Intracellular ATP pool turnover Tile central role of ATP in the stoichiometric coupling of energy-yielding and energy-requiring metabolic reactions has been known since the pioneering research of Lipmann (1941). The steady-state intracellular concentration of ATP in viable microorganisms (prokaryotes and eukaryotes) appears to be well-regulated at a value of 2 to 6 nmol ATP mg ~dry weight (-1 to 3 TriM)regardless of growth rate, culture condition, or mode of nutrition (Karl, 1980). Previous studies of ATP pools in microorganisms
258
and in environmental samples have emphasized the futility of extrapolating these static measurements to estimates of metabolic energy flux. Consequently, it is the turnover rate of the ATP pool, rather than the steady-state concentration of that pool, which varies in proportion to cellular metabolic energy requirements. It follows then, that direct measurements of ATP pool turnover rates, when coupled with independent estimates of ATP pool size, should provide useful information on biological energy flux in cells, populations, or natural microbial assemblages. Cellular ATP pool turnover results from the hydrolysis of one or both of the anhydride-bound PO~ groups (the [3-P and Y-P or 'high-energy' phosphate bonds) via orthophosphate or pyrophosphate cleavage, followed by subsequent regeneration of ATP by either substrate-level, oxidative, or photophosphorylation processes. Because ATP pool turnover defines a cycle, this essential set of metabolic processes results neither in net removal of adenine nucleotide molecules from the intracellular pool nor requires coupled biosynthesis thereof. This is important in the analytical procedures employed to distinguish between the turnover cycles of ATP and total adenine nucleotide (TAN) pools (Karl, 1993). In theory, ATP pool turnover rates could be measured under steadystate metabolic conditions (i.e. d[ATP]/dt = 0) by estimating either the rate of ATP formation or rate of ATP utilization, if either process could be temporarily suspended without affecting the other. Such ATP pool transitions have been observed in microorganisms following the rapid removal of oxygen (for aerobic heterotrophs) or light (for phototrophs), but it is uncertain whether the results derived from these harsh experimental perturbations yield reliable estimates of ATP pool dynamics. Karl and Bossard (1985a) devised a novel method for ATP pool turnover in cell cultures or natural populations of microorganisms. The procedure relies upon the use of :2pi (or ~Pi) as a tracer for P-flux through the acid anhydride-bound P (13-Pand 7-P) groups of cellular ATE The uptake of ':Pi also results in the labeling of the c~-P group of ATE if the population in question is actively growing (i.e. if there is a net removal of adenine nucleotides for biosynthesis). Consequently, in order to uniquely assess the labeling of ATP derived from energy flux, one must be able to separate 13-P and 7-P labeling from c~-P (Karl and Bossard, 1985a). On the other hand, c~-P labeling defines TAN pool turnover and therefore, ~Pi labeling can be used to estimate both ATP and TAN pool turnover rates. The uptake, or salvage, of exogenously added nucleic acid precursors (e.g. adenine, thymidine, uridine) in preference to de m~vo synthesis is a well-documented characteristic of aquatic microbial communities (Karl, 1979; Fuhrman and Azam, 1980). This phenomenon comprises the theoretical basis for the use of l~HIadenine, [~H]thymidine, and I~H]uridine in ecological studies of nucleic acid synthesis. An important aspect of the incorporation of nucleic acid precursors is the observation that the total flux of precursor into the nucteotide triphosphate pools (i.e. ATE TTI; UTP, etc.), which is the result of the combined effects of salvage and de novo synthesis, is in equilibrium with the removal of the triphosphate precursors required for cellular biosynthesis. Furthermore, it is well 259
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documented that the TAN pool (i.e. steady-state intracellular concentration of [ATP + ADP + AMP]) in microorganisms does not vary with changes in the specific growth rate (Karl, 1980; Chapman and Atkinson, 1977). Because cellular biosynthesis is directly related to growth rate, a positive correlation is expected to exist between cellular adenine nucleotide flux and growth. It follows then that the turnover rate of the TAN pool (and most likely, of all ribonucleotide and deoxyribonucleotide pools in general) must vary in direct proportion to the rates of nucleic acid synthesis and hence, net growth. The TAN pool turnover time is defined as the average residence time of a molecule in the intracellular pool before it is removed for macromolecular biosynthesis (i.e. the steady-state pool concentration divided by the steady-state rate of synthesis or removal). If a radioactive precursor such as [~H]adenine is added to a growing culture or natural population of microorganisms, the TAN pool turnover time can be calculated by monitoring the change in the specific activity (SA) of the ATP pool with incubation time. TAN pool turnover rate can also be measured by monitoring the labeling kinetics of the c~-P position of ATP following the addition of ~2Pi (or ~Pi) to a sample (Karl and Bossard, 1985b). Exactly one turnover cycle has been completed when the ATP pool has achieved an SA that is equal to 50% of the value at isotopic equilibrium. Chapman and Atkinson (1977) have estimated TAN pool turnover rates as a function of growth rate for Escherichia coli and SalmoJlella typhimurium. They conclude that the TAN pool 'turns over' (i.e. is completely utilized for biosynthesis and is replenished through salvage and de novo synthesis) 30 to 50 times per generation regardless of generation time. This prediction has been tested using a diverse variety of marine microorganisms (Karl et al., 1987). The results indicate that the TAN pool turnover time is positively correlated with generation time and averages 2.2c~ of the generation time (i.e. TAN pool turns over 45 times per generation). Consequently, a single time-course incubation with ~Pi (~Pi), followed by isolation and selective hydrolysis of the ATP pool, can be used to assess the independent labeling kinetics of the (x-P, [3-P and y-P moieties of ATE. The turnover rate of the c~-P position provides quantitative information on TAN pool turnover and, hence, growth rate, and the turnover rate of the ]3-P and ~/-P positions provides quantitative information on ATP pool turnover and, hence, energy flux. The labeling of ~/-P can also be used to estimate the BAP pool (see following section). Specific ecological applications of the ATP pool and TAN pool turnover for microbial growth rate and energy flux determinations have been presented elsewhere (Karl and Bossard, 1985a,b; Bossard and Karl, 1986; Laws et al., 1986; Karl et al., 1987; Karl, 1993).
Principle: Biologically available P Regardless of the rigor and precision with which P-containing pools are measured, the ecological significance of these analytical determinations will be incomplete until reliable estimates of the BAP pool are routinely
260
available. In addition to Pi, which is generally the preferred substrate for microorganisms, the P contained in a variety of polymeric inorganic compounds, in monomeric and polymeric organic compounds and in selected P-containing minerals is available to some or all microorganisms; indeed some microorganisms may prefer ester-linked P sources to free orthophosphate (Tarapchak and Moll, ]990; Cotner and Wetzel, 1992). However, the bioavailability of most organic-P pools depends criticallv on ambient Pi pool concentrations and on the expression of specific transport, salvage and hydrolytic enzymes. Because many of these enzymes are induced by low Pi, bioavailability may be a variable, time- and habitat condition-dependent parameter, rather than an easily predicted or measured metric. Assessment of the BAP may also depend on the timescale of consideration: substrates that appear to be recalcitrant on short timescales (e.g. < ] day) may readily fuel longer-term (annual to decadal) microbial metabolism. BAP can be estimated using the y-P labeling technique described later for ATP pool turnover. Because the intracellular ATP pool turns over rapidly, the P-specific radioactivity in the y-P position of the P-ATP pool will reach equilibrium not only with the extracellular Pi pool but with the potentially larger pool of extracellular P that is available to the microbial assemblage at that time, the so-called BAP pool. The only difference between the determination of ATP y-P labeling and estinration of BAP is that the latter determination demands a direct measurement of the total P-ATP pool, as well as the amount of ~P (or ~P) contained in the isolated ¥-P fraction thereof. This provides the data necessary to calculate the P-specific radioactivity of the y-P in the intracellular ATP pool (i.e. ~P (or ~P) tool L)which, at isotopic equilibrium (i.e. five turnover cycles) is identical to the specific radioactivity of the extracellular BAP pool.
A T P pool t u r n o v e r and B A P
¢. ,D
m
E q u i p m e n t and reagents
U
• Acid-cleaned polycarbonate incubation bottles (0.1 to 4 I, depending on experimental design) • On-deck temperature- and light-controlled incubator, or in situ incubation array • Filtration gear, heating block and other ATP extraction and detection equipment and supplies • Vacuum evaporator (Savant Speed-Vac or equivalent) • Microcentrifuge and gyrotatory shaker cable • Thin layer chromatography (TLC) glass tanks with covers • TLC plates (20 × 20 cm), polyethyleneimine (PEI) impregnated cellulose (Polygram CEL 300 PEI, Machery-Nagel, Fisher #NC-9254648) • High-intensity ultraviolet (UV) lamp • Radiolabeled orthophosphate (32P©4 or 3?©4) as a stock solution of approximately 18.5 MBq ml' (500 pCi ml ') (ICN Radiochemicals)
261
u~ O •
O a.
• ATP (0.5 mM in Tris buffer 20 mM, pH 7.4, Sigma #A-6144) • Formic acid (0.2 M, Baker #1-0128) • Potassium phosphate solution (0.85 M, pH 3.4, Fisher #P288) • Magnesium chloride (0.35 M, Fisher #M87) • Activated charcoal slurry: 25 mg charcoal per ml of 0. I M H3PO 4 (activated charcoal, Sigma #C-4386) • ATPase, porcine cerebral cortex (EC 3.6'. 1.3; Sigma Chemical Co, #A7510) prepared in distilled deionized water ( D D W ) to a concentration of I unit ml ' and stored frozen a t - 2 0 ° C until needed. Alternatively apyrase (Sigma Chemical Co, #A6132) can be used (10 units m ' in D D W ) and stored frozen
•
Liquid scintillation counter (LSC)
Assay
1. Incubate freshly collected seawater samples under defined temperature and light conditions in the presence of exogenous 32pi (or 3~pi); typically 20-40 MBq 1 '. 2. At predetermined incubation periods, subsamples are removed and particulate ATP pools are extracted in boiling Tris buffer and stored frozen. 3. Thaw samples; centrifuge at 1000g for 15 min to remove filter debris. 4. Transfer 3.5 to 4.0 ml of the supernatant to clean scintillation vials. 5. Evaporate to dryness in vacuo (Speed Vac). The samples are then reconstituted in 100-200 pl DDW. 6. In clean disposable glass tube mix 10 pl of concentrated ATP extract with 10 pl of non-radioactive ATP (0.5 mM) solution. Spot 10 pl of this mixture on predetermined location approximately 1 cm from bottom edge on a PEI plate. About 8-10 lanes can be prepared per plate. 7. Air-dry plate and then wash twice (5 min each) by immersion in DDW (1 1). Allow plate to dry (this step can be accelerated by using a hairdryer). 8. Develop plate in formic acid (0.2 M)' to top of plate in a closed TLC chamber. 9. Remove and immediately immerse in DDW (1 1) for 5 min. Allow to dry. 10. Develop plate in the same direction in potassium phosphate solution (0.85 M, pH 3.4) to approximately 2 cm from top edge of plate in closed TLC chamber. 11. Wash plate (2 x 5 min, 1 1 DDW) and air dry. 12. The ATP spots are visualized under UV-light, circled with a pencil and cut out with a pair of scissors. Recovery is typically >90%. 13. The excised PEI cuttings are placed into scintillation vials containing I m! of 0.35 M MgC12 and placed on shaker table for I h to elute ATP from the PEI matrix.
262
14. A 101Jl aliquot from the elute is diluted into l m l Tris buffer (20 mM, pH 7.4) and assayed for total ATP by the firefly bioluminescence reaction. This is to determine the recovery of the ATP initially applied to the PEI plate. 15. To perform the ATPase hydrolysis reaction place 7501.ll of the 'TLC'-purified ATP solution (in 0.35 M MgC12) into a 12 x 75 mm glass culture tube containing; add 250 Ill DDW, 10 1~1 KC1 (1 M), 20 lJl NaC1 (5 M) and 25 Ill AMP (3 mM). 16. Add 501Jl of the ATPase solution (stock at 1 unit ml '). Mix thoroughly. 17. Immediately remove 10 1_ll of mixture and dilute into 1 ml Tris (20 mM, pH 7.4) to quench the reaction and assay for 'time zero' ATP (as above for ATP determination). 18. The ATP procedure is repeated (10 lJl sample, as above) after 10 to 15 min with one or two representative samples to establish the initial rate of hydrolysis of ATP to ADP. Once the rate is known the time for complete hydrolysis can be calculated (normally 2040 min). The rate of hydrolysis is expected to be similar, however, complete ATP hydrolysis should be confirmed for each sample before proceeding with the separation and purification of products. 19. Remove 250 lJ1 of the sample and place in scintillation vial for LSC counting of the 'total radioactivity' (i.e. o% [3- and ~_32pof ATP). 20. Place a second 500 lJl aliquot into a 1.5 ml microcentrifuge tube containing 500 Ill of the charcoal slurry. Mix thoroughly (vortex). 21. Centrifuge at 13 000g for 10 min. 22. Remove 750 1,11of supernatant is placed into a scintillation vial for LSC counting of the y-32p-activity. 23. Calculate the specific radioactivity of the ATP pool over time. 24. The specific y-3'P-activity of the intracellular ATP pool when it has reached its isotopic equilibrium (i.e. maximum) corresponds to the specific activity of the precursor pool (i.e. the bioavailable P (BAP) pool) and the size of the BAP pool can be determined. Note: If apyrase is used instead of ATPase in step #15; place 750 pl of the 'TLC'-purified ATP solution (in 0.35 M MgCI2) into a 12 x 75 mm glass culture tube containing; 250 1,11DDW, 20 lal CaC12 (1 M). Add 25 1.11 apyrase (10 units ml ') and mix thoroughly. Proceed from step 21. The apyrase cleaves both the 13- and ,},_32pof ATP and this has to be taken into account when calculating the specific activity of the intracellular ATP pool and in estimating the size of the BAP pool. Calculations: From the data obtained above, the turnover time of the total ATP (13-and ,yg2p labeling) and or total adenine (TAN) pool (o~-32p labeling) can be calculated and from that energy flux and community growth rate respectively. The change in the ATP or TAN pool-specific activity (SA; nCi pmol-~), predicted by radiotracer theory, follows an exponential function of the incubation time. The decimal equivalent of SA at any time (SA,) can be described by the equation: SA, = 1 - (2 "),
263
¢, m
O3 U
O e-
where N is the n u m b e r of t u r n o v e r cycles o b s e r v e d during the incubation period. ATP or TAN pool turnover time (T) can be calculated from the expression: T -- t / N . At incubation times _> 5 the pools are in isotopic equilibrium and would, in theory, not change until the exogenous precursor source is exhausted, and pool delabeling begins. Once T has been d e t e r m i n e d it can be used to extrapolate energy flux (EF) and specific g r o w t h rate: EF (kcal 1 ' h ') = -22 ([ATP]/T,~.~,), w h e r e [ATP] is equal to the total particulate ATP pool (M) and T.,r~is ATP pool turnover time (h). To estimate growth rate, the TAN pool t u r n o v e r time is a s s u m e d to be equivalent to 2-3% of the generation time (i.e. the doubling time (T~) is on average 45 x T,,,~).
References Ammerman, J. W. and Azam, F. (1981). Dissolved cyclic adenosine monophosphate (cAMP) in the sea and uptake of cAMP by marine bacteria. Mar. Ecol. Prog. Set. 5, 85-89. Ammerman, J. W. and Azam, E (1985). Bacterial 5'-nucleotidase in aquatic ecosystems: a novel mechanism of phosphorus regeneration. Science 227, 1338-1340. Armstrong, E A., Williams, P. M. and Strickland, J. D. H. (1966). Photooxidation of organic matter in seawater by ultraviolet radiation, analytical and other applications. Nature 211, 481-483. Atkinson, M. J. (1987). Rates of phosphate uptake by coral reef flat communities. Limnol. Oceanogr. 32, 426-435. Azam, F. and Hodson, R. E. (1977). Dissolved ATP in the sea and its utilisation by marine bacteria. Nature 267, 696-698. Benitez-Nelson, C. R. and Buesseler, K. O. (1999). Variability of inorganic and organic phosphorus turnover rates in the coastal ocean. Nature 398, 502-505. Bj6rkman, K. and Karl, D. M. (1994). Bioavailability of inorganic and organic phosphorus compounds to natural assemblages of microorganisms in Hawaiian coastal waters. Mar. Ecol. Pro~. Ser. 111, 265-273. Bj6rkman, K. and Karl, D. M. (2001). A novel method for the measurement of dissolved nucleotides in seawater: Applications in marine microbial ecology. Submitted to Journal of Microbiological Methods. Bj6rkman, K., Thomson-Bulldis, A. L. and Karl, D. M. (2000). Phosphorus dynamics in the North Pacific subtropical gyre. Aquatic Microbial. Ecol., 22, 185-198. Bossard, P. and Karl, D. M. (1986). The direct measurement of ATP and adenine nucleotide pool turnover in microorganisms: A new method for environmental assessment of metabolism, energy flux and phosphorus dynamics. J. Plankton Res. 8, 1-13. Brown, E. J., Harris, R. F. and Koonce, J. E (1978). Kinetics of phosphate uptake by aquatic microorganisms: Deviations from a simple Michaelis-Menten equation. LimJlol. Oceano~r. 23, 26-34. Carlucci, A. E and Silbernagel, S. B. (1966). Bioassay of seawater. I. HC-uptake method for the determination of concentrations of vitamin B,: in seawater. Can. J. Microbiol. 12, 175-183. Cerebella, A. D. and Antia, N. J. (1986). The determination of phosphonates in seawater by fractionation of the total phosphorus. Mar. Chem. 19, 205-210.
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Cembella, A. D., Antia, N. J. and Taylor, E J. R. (1986). The determination of total phosphorus in seawater by nitrate oxidation of the organic component. Water Res. 20, 1197-1199. Chamberlain, W. and Shapiro, J. (1969). On the biological significance of phosphate analysis; comparison of standard and new methods with a bioassay. Limnol. Oceanogr. 14, 921-927. Chapman, A. G. and Atkinson, D. E. (1977). Adenine nucleotide concentrations and turnover rates. Their correlation with biological activity in bacteria and yeast. Adv. Microb. Physiol. 15, 253 306. Clark, L. L., lngall, E. D. and Benner, R. (1999). Marine organic phosphorus cycling: Novel insights from nuclear magnetic resonance. Amer. J. Sci. 2999, 724-737. Correll, D. L. (1965). Pelagic phosphorus metabolism in Antarctic waters. Limnol. Oceanogr. 10, 364 370. Cotner, J. B. and WetzeI, R. G. (1992). Uptake of dissolved inorganic and organic phosphorus compounds by phytoplankton and bacterioplankton. Limnol. Oceauogr. 37, 232-243. Cuhel, R. L., Jannasch, H. W., Taylor, C. D. and Lean, D. R. S. (1983). Microbial growth and macromolecular synthesis in the northwestern Atlantic Ocean. Limnol. Oceano~r. 28, 1-18. deDuve, C. (1991). Blueprint for ~ Cell: Tile Nature and Ori~4in of Life. Nell Patterson Publishers, Burlington, NC. DeFlaun, M. E, Paul, J. H. and Davis, D. (1986). Simplified method for dissolved DNA determination in aquatic environments. Appl. EJrviron. Microbiol. 52, 654-659. Fernandez, J. A., Niell, E X. and Lucena, J. (1985). A rapid and sensitive automated determination of phosphate in natural waters. Limuol. Oceano~r. 30, 227-230. Findlay, R. H., King, G. M. and Watling, L. (1989). Efficacy of phospholipid analysis in determining microbial biomass in sediments. Appl. EnviJvH. Microbiol. 55, 2888-2893. Fiske, C. H. and Subbarow, Y. (1925). The colorimetric determination of phosphorus. ]. Biol. Chem. 66, 375-400. Francko, D. A. (1984). Relationships between phosphorus functional classes and alkaline phosphatase activity in reservoir lakes. J. Freshwater Ecol. 2, 541-547. Fuhrman, J. A. and Azam, F. (1980). Bacterioplankton secondary production estimates for coastal waters of British Columbia, Antarctica and California. Appl. EHviro11. Microbiol. 39, 1085-1095. Harrison, W. G. (1983). Uptake and recycling of soluble reactive phosphorus by marine microplankton. Mar. Ecol. Pro~. Set. 10, 127 135. Harrison, W. G., Azam, F., Renger, E. H. and Eppley, R. W. (1977). Some experiments on phosphate assimilation by coastal marine plankton. Mnr. Biol. 40, 9-18. Holm-Hansen, O. and Booth, C. R. (1966). The measurement of adenosine triphosphate in the ocean and its ecological significance. LimHol. Ocealzogr. 11, 510-519. Holm-Hansen, O., Sutcliffe, W. H., Jr. and Sharp, J. (1968). Measurement of deoxyribonucleic acid in the ocean and its ecological significance. Limnol. Occanogr. 13, 507-514. Hoppe, H.-G. (1993). Use of fluorogenic model substrates for extracellular enzyme activity (EEA) measurement of bacteria. In: Handbook of Methods in Aquatic Microbial Ecology (P. F. Kemp, B. F. Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 423-431. Lewis Publishers, Boca Raton, FL. Hudson, J. J. and Tayk)r, W. D. (1996). Measuring regeneration of dissolved phosphorus in planktonic communities. Limllol. Oceano~¢r. 41, 1560-1565.
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Ingall, E. D., Schroeder, P. A. and Berner, R. A. (1990). The nature of organic phosphorus in marine sediments: New insights from ~P NMR. Geochim. Cosmochim. Acta 54, 2617-2620. Johannes, R. E. (1964) Uptake and release of dissolved organic phosphorus by representatives of a coastal marine ecosystem. Limnol. Oceanogr. 9, 224-234. Jones, P. G. W. and Spencer, C. P. (1963). Comparison of several methods of determining inorganic phosphate in seawater. J. Mar. Biol. Assoc. UK 43, 251-273. Karl, D. M. (1978a). A rapid sensitive method for the measurement of guanine ribonucleotides in bacterial and environmental extracts. Anal. Biochem. 89, 581-595. Karl, D. M. (1978b). Occurrence and ecological significance of GTP in the ocean and in microbial cells. Appl. Environ. Microbiol. 36, 349-355. Karl, D. M. (1979). Adenosine triphosphate and guanosine triphosphate determinations in intertidal sediments. In: Methodology for Biomass Determinations and Microbial Activities in Sediments (C. D. Litchfield and P. L. Seyfried, Eds) ASTM STP 673, pp. 5 20. Amer. Soc. Testing and Materials, Philadelphia. Karl, D. M. (1980). Cellular nucleotide measurements and applications in microbiaI ecology. Microbiol. Rev. 44, 739-796. Karl, D. M. (1981). Simultaneous rates of ribonucleic acid and deoxyribonucleic acid syntheses for estimating growth and cell division of aquatic microbial communities. Appl. Environ. Microbiol. 42, 802-810. Karl, D. M. (1993). Adenosine triphosphate (ATP) and total adenine nucleotide (TAN) pool turnover rates as measures of energy flux and specific growth rate in natural populations of microorganisms. In: Currellt Methods in Aquatic Microbial Ecology (P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds) pp. 483-494. Lewis Publishers, Boca Raton, FL. Karl, D. M. and Bailiff, M. D. (1989). The measurement and distribution of dissolved nucleic acids in aquatic environments. Limnol. Oceanogr. 34, 543-558. Karl, D. M. and Bossard, P. (1985a). Measurement and significance of ATP and adenine nucleotide pool turnover in microbial cells and environmental samples. J. Microbiol. Methods 3, 125-139. Karl, D. M. and Bossard, P. (1985b). Measurement of microbial nucleic acid synthesis and specific growth rate by ~PO~ and [~H]adenine: Field comparison. Appl. Environ. Microbiol. 50, 706 709. Karl, D. M. and Dobbs, E C. (1998). Molecular approaches to microbial biomass estimation in the sea. In: Molecular Approaches to the Study of the Ocean (K. E. Cooksey, Ed.), pp. 29-89. Chapman & Hall, London. Karl, D. M. and Holm-Hansen, O. (1978). ATP, ADP and AMP determinations in water samples and algal cultures. In: Handbook of Phycological Methods, Vol. III. Physiological and Biochemical Methods (J. A. Hellebust and J. S. Craigie, Eds), pp. 197-206. Cambridge University Press. Karl, D. M. and Tien, G. (1992). MAGIC: A sensitive and precise method for measuring dissolved phosphorus in aquatic environments. Limnol. Oceanogr. 37, 105-116. Karl, D. M. and Tien, G. (1997). Temporal variability in dissolved phosphorus concentrations in the subtropical North Pacific Ocean. Mar. Chem. 56, 77-96. Karl, D. M. and Yanagi, K. (1997). Partial characterization of the dissolved organic phosphorus pool in the oligotrophic North Pacific Ocean. Limnol. Oceanogr. 42, 1398-1405. Karl, D. M., Jones, D. R., Novitsky, J. A., Winn, C. D. and Bossard, P. (1987). Specific growth rates of natural microbial communities measured by adenine nucleotide pool turnover. J. Microbiol. Methods 6, 221-235. Karl, D. M., Dore, J. E., HebeI, D. V. and Winn, C. (1991). Procedures for particulate
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carbon, nitrogen, phosphorus and total mass analyses used in the US-JGOFS Hawaii Ocean Time-Series Program. In: Marine Particles: Analysis and CharacterizatiolJ (D. Spencer and D. Hurd, Eds) Geophysical Monograph 63, pp. 71-77. American Geophysical Union. Karl, D. M., Christian, J. R., Dore, J. E., Hebel, D. V., Letelier, R. M., Tupas, L. M. and Winn, C. D. (1996). Seasonal and interannual variability in primary production and particle flux at Station ALOHA. Deep Sea Res. II 43, 539-568. Knauer, G. A., Martin, J. H. and Bruland, K. W. (1979). Fluxes of particulate carbon, nitrogen and phosphorus in the upper water column of the northeast Pacific. Deep-Sea Res. 26, 97-108. Krempin, D. W., McGrath, S. M., SooHoo, J. B. and Sullivan, C. W. (1981). Orthophosphate uptake by phytoplankton and bacterioplankton from the Los Angeles Harbor and southern California coastal waters. Mar. Biol. 64, 23-33. Kuenzler, E. J. and Ketchum B. H. (1962). Rate of phosphorus uptake by Phaeodactylum tricormttum. Biol. Bull. 123, 134-145. Kuenzler, E. J. and Perras, J. P. (1965). Phosphatases of marine algae. Biol. Bull. 128, 271-284. Lal, D. and Lee, T. (1988). Cosmogenic ~2P and '~P used as tracers to study phosphorus recycling in the ocean. Natulv 333, 752-754. Lal, D., Chung, Y., Platt, T. and Lee, T. (1988). Twin cosmogenic radiotracer studies of phosphorus cycling and chemical fluxes in the upper ocean. Limnol. Ocealwgr. 33, 1559-1567. Laws, E. A., Jones, D. and Karl, D. M. (1986). Method for assessing heterogeneity in turnover rates within microbial communities. Appl. Em~iro,. Microbiol. 52, 866-874. Lemasson, L. and Pages, J. (1981). Excretion of dissolved organic phosphorus in tropical brackish waters. Est. Coastal Shelf Sci. 12, 511-523. Levin, G. B., Clendenning, J. R., Chappelle, E. W. et al. (1964). A rapid method for detection of microorganisms using the ATP assay. BioScience 14, 37-38. Lin, R. I. S. and Schjeide, O. A. (1969). Micro estimation of RNA by the cupric ion catalyzed orcinol reaction. Anal. Biochem. 27, 473-483. Lipmann, E (1941). Metabolic generation and utilization of phosphate bond energy. Adv. Enzymol. Relat. Areas Mol. Biol. 1, 99-162. Maeda, M. and Taga, N. (1973). Deoxyribonuclease activity in seawater and sediment. Mar. Biol. 20, 58-63. Maki, J. S., Sierszen, M. E. and Remsen, C. C. (1983). Measurement of dissolved adenosine triphosphate in Lake Michigan. Can. J. Fish. Aquatic Sci. 40, 542-547. McGrath, S. M. and Sullivan, C. W. (1981). Community metabolism of adenylates by microheterotrophs from the Los Angeles and Southern California coastal waters. Mar. Biol. 62, 217-226. Menzel, D. W. and Corwin, N. (1965). The measurement of total phosphorus in seawater based on the liberation of organically bound fractions by persulfate oxidation. Limnol. Ocealtogr. 10, 280-282. Monaghan, E. J. and Ruttenberg, K. C. (1999). Dissolved organic phosphorus in the coastal ocean: Reassessment of available methods and seasonal phosphorus profiles from the Eel River Shelf. Limnol. Oceanogr. 44, 1702-1714. Murphy, J. and Riley, J. P. (1962). A modified single solution method for the deterruination of phosphate in natural waters. Anal. Chim. Acta 27, 31-36. Nawrocki, M. P. and Karl, D. M. (1989). Dissolved ATP turnover in the Bransfield Strait, Antarctica during a spring bloom. Mar. Ecol. Prog. Set. 57, 35-44. Ormaza-GonzMez, F. I. and Statham, P. J. (1991). Determination of dissolved inorganic phosphorus in natural waters at nanomolar concentrations using a long capillary cell detector. Aizal. Chim. Acta 244, 63-70.
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Tarapchak, S. J. and Moll, R. A. (1990). Phosphorus sources for phytoplankton and bacteria in Lake Michigan. J. Plankton Res. 12, 743-758. Thomson-Bulldis, A. and Karl, D. M. (1998). Application of a novel method for phosphorus determinations in the oligotrophic North Pacific Ocean. Limnol. Oceanogr. 43, 1565-1577. Waser, N. A. D., Bacon, M. P. and Michaels, A. E (1996). Natural activities of ~2pand ~P and the ~p/~2p ratio in suspended particulate matter and plankton in the Sargasso Sea. Deep-Sea Res. Ii 43, 421-436. Watt, W. D. and Hayes, E R. (1963). Tracer study of the phosphorus cycle in sea water. Limnol. Oceanogr. 8, 276-285. White, D. C., Bobbie, R. J., Morrison, S. J., Oosterhof, D. K., Taylor, C. W. and Meeter, D. A. (1977). Determination of microbial activity of estuarine detritus by relative rates of lipid biosynthesis. Limnol. Oceanogr. 22, 1089-1099. White, D. C., Davis, W. M., Nickels, J. S., King, J. D. and Bobbie, R. J. (1979). Determination of the sedimentary microbial biomass by extractible lipid phosphate. Oecologia 40, 51-62. White, R. H. and Miller, S. L. (1976). Inositol isomers: occurrence in marine sediments. Science 193, 885-886.
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14 Nitrogen Fixation: Nitrogenase Genes and Gene Expression JP Zehr and PJ Turner Department of Ocean Sciences,University of California-Santa Cruz, Santa Cruz, California 95064, USA
CONTENTS Introduction Methods overview PCR amplification of nitrogenase genes and phylogenetic analysis Primers and controls for PCR Genomic DNA extraction Polymerase chain reaction (PCR) Nitrogenase gene expression Reverse transcriptase-polymerase chain reaction (RT-PCR) Problems, limitations and caveats Applications Future directions and alternative approaches
INTRODUCTION Biological nitrogen fixation is the enzymatic reduction of atmospheric dinitrogen to ammonium. This process, a key component of the nitrogen cycle, is important in many ecosystems when biologically more available forms, such as nitrate or ammonium, are present in small amounts relative to biological demand for growth. The capability for nitrogen fixation is widely dispersed among prokaryotic taxa. Very divergent, distantly related organisms are able to fix nitrogen. On the other hand, not all taxa within a specific group are able to fix nitrogen. For example, there are nitrogen-fixing and non-nitrogen-fixing species of unicellular cyanobacteria. The nitrogenase enzyme is a multi-component enzyme that typically consists of the iron (Fe) protein and the molybdenum iron (MoFe) protein (Howard and Rees, 1996). This nitrogenase is termed 'conventional'. Alternative forms of the protein exist that replace Mo with vanadium ('alternative') or Fe ('second alternative'). The conventional nitrogenase is encoded by the nifHDK genes, which are often found in contiguous arrangement within the genome. Alternative nitrogenases (alternative and second alternative) also contain nifH, but contain a third protein in the counterpart to the Mo protein, which is encoded by nifG (nifDGK). The METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0-12-521530-4
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nifH genes in all of these nitrogenases are highly conserved (Howard and Rees, 1996). The nitrogenases share a small, but significant, degree of similarity with chlorophyllide reductases and with nifH-like genes that are found in some Archaea (methanogens). The many different types of diazotrophic prokaryotic taxa make it difficult to determine which microorganism is responsible for observed nitrogen fixation rates, or to determine the potential for microbial communities to respond to nitrogen fixation conditions. A genetic approach for assessing the composition of diazotrophic communities and for assaying the extent of gene expression can be used as a powerful complementary tool for tracer and analogue (acetylene reduction) assays for measuring nitrogen fixation rates. These approaches provide for a comprehensive portrait of nitrogen fixation in natural assemblages of microbes.
METHODS OVERVIEW Nitrogenase genes can be detected and characterized by amplification from environmental samples using the polymerase chain reaction (PCR). Amplification of nitrogenase genes indicates that nitrogen-fixing microorganisms are present, but not whether or not they are actively fixing nitrogen. By coupling the PCR assay with reverse transcription (RT-PCR) microorganisms that are actively expressing the nitrogenase enzyme can be detected. Once genes are amplified, the diversity of sequences can be determined by a number of means, including cloning and sequencing of individual amplification products. The amplification products can potentially be quantified, using a recombinant competitive template that is amplified by the niflq primers, but which differs in size, allowing the nifH product and the competitor product to be distinguished by gel electrophoresis (Zimmermann and Mannhalter, 1996; Larrick, 1997). The competitive PCR approach will not be covered here. The amplification product can be cloned, and individual clones sequenced to obtain the nifH gene sequences of individual target molecules in the PCR or RT-PCR reaction. In this discussion, we will describe (1) the primers used for nifH amplification, (2) the methods used to extract genomic DNA and mRNA, (3) the alignment and analysis of nifH sequences and (4) the RT-PCR protocol (Figure 14.1).
P C R A M P L I F I C A T I O N OF N I T R O G E N A S E GENES A N D P H Y L O G E N E T I C A N A L Y S I S Principle There are highly conserved regions in the nifH protein amino acid sequences, which can be used to design degenerate oligonucleotide primers. DNA samples are prepared to reduce inhibition of amplification as much as possible. The use of a nested PCR and RT-PCR technique greatly 272
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Figure 14.1. Overview of methods for amplifying and characterizing nifDNA and RNA, Nucleic acid samples can be amplified directly with PCR, or reverse transcribed to cDNA with reverse-transcriptase prior to PCR in the RT-ECR technique. Amplification products can be quantified by a variety of approaches, including competitive PCR. The amplification products can be cloned to create a llifH library from the sample, and the clones screened and analyzed by phylogenetic analysis.
reduces the effects of inhibition. Very small samples can be assayed, thereby adding only minimal amounts of associated contaminants and inhibitors, while still retaining a high degree of sensitivity. The D N A or d e d u c e d amino acid sequences of nifH obtained from amplification, cloning and sequencing can be used for phylogenetic analysis. In general, there are slight differences between trees based on D N A sequences, and minimal differences between trees based on amino acid sequences and sequences based on DNA sequences without the third base of the codons (Zehr el al., 1997). The resulting tree distinguishes between the major nifH gene types (alternative, nifH-like, etc.), and can be used as a f r a m e w o r k to identify nitrogen-fixing microorganisms on the basis of sequences obtained directly from the environment via PCR or RTPCR (Figure 14.2).
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PRIMERS A N D C O N T R O L S
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Figure 14.2. Phylogenetic tree of nifH sequences showing major nifH clusters. The tree was constructed by a distance method using the program TREECON. Chlorophyllide reductase gene sequences were used as the outgroup for the tree, and are not shown on the tree. The triangles denote major clades. Bootstrap values for 100 replicates are indicated at nodes, and indicate the number of times the sequences to the right of the node group together in the analysis. The question mark highlights a deeply branching clade recently found in aquatic environments, and demonstrates that the identification of new deeply divergent sequences can be problematic and emphasizes the need to continue to obtain representative nifH sequences from cultivated microorganisms.
1995) (Table 14.1). Two sets of these primers target virtually identical regions (Zehr and McReynolds, 1989; O h k u m a et al., 1996); some include a 5' restriction site to facilitate cloning (Ohkuma et al., 1996; Ben-Porath et al., 1993). We have found that the addition of a restriction site in the oligonucleotide primers (primers used by Ben-Porath et al., 1993) can reduce amplification efficiency and m a y have implications for selectivity. We have also found that the effectiveness of the degenerate primers is very sensitive to the procedures used to purify the primers. We have had very good success with primers that are PAGE and HPLC purified (synthesized by N e w England Biolabs, Inc.). A previously cloned nifH fragment is used as a positive control. The cloned fragment should include regions c o m p l e m e n t a r y to all pairs of primer sequences used, such as nested primer sets.
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Equipment Basic filtration e q u i p m e n t is needed for preparing samples including a v a c u u m apparatus and filtration tower, and low D N A / R N A binding 274
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membrane filters (e.g. 0.2 gm pore size Supor, Gelman or Durapore, Millipore). Also needed is an autoclave, a water bath set to 37°C, a microcentrifuge, a Speedvac system for drying samples, electrophoresis power supply and gel apparatus, 1.5 ml plastic microcentrifuge tubes, 15 ml sterile plastic capped centrifuge tubes, small plastic Whirl-pack bags (4 oz), adjustable automatic pipettes and sterile pipette tips, latex gloves, and PCR cabinets (enclosed clean workspaces with UV light). The PCR cabinets are particularly helpful for reducing contamination problems.
Reagents Sterile procedures must be used throughout and all glassware and tubes should be autoclaved. All reagents should be made with autoclaved and UV-treated water (exposed to 254 nm UV for 20 rain at a distance of 12 cm from the source). Buffer solutions should be prepared in advance, autoclaved and UVtreated when possible (SDS, ammonium acetate, TE). Enzyme solutions should be prepared with pre-treated water and should not be autoclaved or UV-treated after the addition of the enzyme (lysozyme and proteinase K). Phenol, chloroform and ethanol should not be autoclaved. Buffers and reagents needed are NST buffer (400 mM NaC1, 0.75 mM sucrose, 50 mM Tris-HC1, pH 9.0), lysozyme (50 gg gl '), Proteinase K (20 mg ml '), 10% sodium dodecyl sulfate (SDS), 10 M ammonium acetate, TE buffer (10 mM Tris, pH 8.0, 1 mM EDTA), 1:1 phenol chloroform, chloroform, 100% ethanol, 70% ethanol, 0.1 N HC1 and 1.5% agarose.
Method The technique described here is used for amplification from free-living bacteria and cyanobacteria from water samples. The filter and filtration apparatus should be rinsed prior to filtration with a few milliliters of 0.1 N HC1 to prevent contamination with DNA. For densely populated water samples, pre-filtering through a coarse filter (5-50 gm pore size filter or netting) may be necessary prior to the final filtration. A measured quantity of water is filtered onto 0.2 gm filters to collect the cells. Different diameter filters can be used, depending on the concentration of cells and detritus in the water sample, but 47 mm diameter filters allow a 500-2000 ml sample to be filtered in a reasonable amount of time. One liter samples would be typical for extracting DNA from oligotrophic oceans, and a few hundred milliliters at most from estuarine waters. Following filtration, the filter is placed in a Whirl-pack bag and 2 ml NST buffer added. Lysozyme is then added (100 ~tl of 50 gg ~1 ') and the filter agitated or rubbed to solubilize particles on the filter. The filter is incubated in the bag for 30 min in a 37°C water bath. SDS (200 ~1 of a 10% w / v stock) and 50 ~1 of proteinase K (20 mg ml ~) are then added and the bag incubated for 2 h or overnight in a 37°C water bath. The filter is discarded and 500 ~tl of the lysate added to a sterile microcentrifuge tube. The remaining liquid is frozen in a 15 ml tube for future use. Phenol/chloroform (300 ~,L1of 1 : 1 solution) is added
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and the tube vortexed. The sample is then centrifuged for 4 min at 13 000 rpm (at room temperature), the aqueous layer transferred to a new tube, and 300 [11 chloroform added. After vortexing, the sample is centrifuged again for 4 min at 13 000 rpm and the aqueous layer again transferred to a new tube. Ammonium acetate (100 [11 of 10 M stock) is added, the tube filled with 100% cold ethanol and the DNA precipitated overnight at 4°C (or for 1 h at -20°C). During the subsequent washing steps the tubes are oriented in the centrifuge to maintain the same location of the pellet for each spin. The samples are centrifuged for 30 min at 13 000 rpm and the supernatant aspirated carefully, without disturbing the pellet. The pellet is washed with 500 [11 of cold 70c/~ ethanol and centrifuged again for 10 min. Finally, the supernatant is drawn off carefully and the pellet dried using a SpeedVac on low heat. The pellet is resuspended in 50 ~1 TE (pH 8.0) and 10 [11of the sample run on a 1.5% agarose gel to confirm recovery of DNA, and that DNA is of high quality (high molecular weight and lacking a smear of degradation products).
POLYMERASE C H A I N R E A C T I O N Equipment General laboratory equipment as described above, plus a PCR thermal cycler, electrophoresis power supply and gel box, photodocumentation system (Bio-Rad Gel Doc), 0.2 ml PCR tubes, aerosol-free pipette tips.
Reagents Reagents needed are 25 mM MgCI~ (Promega), 10x PCR buffer (Promega), 10 mM each dNTPs (Promega), primers (100 [1M stocks), Taq polymerase (Promega), and 1.5~ agarose.
Method Typically, less than 100 ng of genomic DNA is used in the PCR reaction. Following the protocol described here, there will be adequate DNA for many amplifications from a single genomic DNA preparation, although this will have to be determined empirically for each sample type. Usually higher concentrations of DNA inhibit the reaction, probably because of inhibitors that are not removed by the extraction procedure used above. Often, the genomic DNA recovered is less than can be visualized by gel electrophoresis, but is still sufficient to amplify nitrogenase genes except from very oligotrophic waters. Usually about 1 [11of tile genomic DNA (or dilution of genomic DNA) is added to the PCR reaction. The PCR can be performed as a single amplification with one pair of primers, or as a nested PCR, using two separate sets of primers in successive amplifications. The single-stage PCR is adequate when working with samples with abundant nitrogen-fixers, but the nested PCR provides greater sensitivity.
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The first stage of the nested PCR is performed using nif primers nifH4 and nifH3 (Table 14.1). To prevent the possibility of contamination, the addition of reagents and DNA should be p e r f o r m e d in separate P C R / U V work stations, such as The Clean Spot available through Coy Laboratory Products. All PCR tubes, pipettors, tips, and water should be UV treated (254 n m at a distance of 12 cm from the source) for 20 min prior to setting up the reactions. Pipettes should be dedicated to preparing the core only or for the addition of DNA only. DNA should never be introduced into the core preparation station. While the supplies are being UV treated, the MgCL (25 raM, Promega) and 10x PCR buffer (Promega) should be w a r m e d to 37 ° C. Core mixes m a y be made simultaneously for each round of the nested PCR. For each core (per sample) add 8 Jal of 25 mM MgC12, 5/21 of buffer, 1 ~1 dNTPs (Promega), 0.5 ~tl of the first round primer sets (core 1) and the nested primer sets (core 2) and 33.5 lal of UV-treated Milli-Q water. The concentration of MgCI~ may have to be determined empirically, d e p e n d i n g on the nature of the sample, and even the supplier of the oligonucleotide primers. These quantities are multiplied by the n u m b e r of desired reactions (adding 1 or 2 reactions per core to ensure adequate volu m e for all reactions) and UV treated as described above. Taq polymerase (2.5 U per reaction) is added following the UV treatment. The core reagents are mixed well and 49/dl added to each of the PCR tubes. We have found that contamination is problematic w h e n working with samples with low numbers of genes, w h e n cloning activities are being performed simultaneously in the laboratory, and w h e n a cloned positive control is used. In order to avoid contamination, only one tube at a time from the first round core is m o v e d to the next station for the addition of sample DNA. it is advisable to remove both the sample DNA and the loaded core to a third station before repeating the process with the next tube. This separation of activities, although tedious, reduces cross-tube contamination, particularly from the positive control and competitive templates used in the analyses. We use separate PCR UV hoods for each station. The first stage of the polymerase chain reaction is performed for 25-30 cycles of denaturation at 94°C (1 min), annealing at 57°C (1 rain), and extension at 72°C (1 min) using the nifH4 and nifH3 primers. The reactions are carried out in 50 ~tl volumes. In the nested PCR, 1 gl of the first
Figure 14.3. Amplification of n(]CH mRNA using the nested RT-PCR method. Interpretation of RT-PCR results is dependent upon eliminating the possibility that the amplification products are derived from DNA rather than RNA. Two controls that can be used are (A) amplification of the RNA sample with a normal PCR and (B) amplification by RT-PCR after RNasing the sample. (A) PCR controls for RT-PCR. Lane 1: RNA positive control included with each sample run. The positive control is RNA extracted from a nitrogen-fixing microorganism. Lane 2: The sample amplified with PCR, without the RT step. This lane should appear negative if the RNA sample is not contaminated with DNA. Lane 3: A positive nifH DNA control to show that the PCR conditions were adequate for amplification of DNA. (B) RNased controls. Lane 1: RNA sample treated with RNase. lane 2: RNA sample amplified with RT-PCR. Lane 3: Negative control for complete RT-PCR procedure. Lane 4: uifH DNA positive control to test for correct amplification. Lane 5: Negative control for PCR amplification. 278
round PCR product is added to the second reaction, which is set up identical to the first reaction except for adding nifH1 and nifH2 primers.
Cloning Tile amplified fragments (approximately 359 base pairs, see Figure 14.3) are gel purified and cloned into a plasmid 'T' vector (such as pTYBlue; Novagen, Madison, Wis. or pGEM T vector, Promega) to create a clone
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library. Burlage (1998) provides a good background discussion of general cloning techniques. Cloning kits for cloning of PCR fragments are available from a number of manufacturers. Using blue-white selection, colonies are picked, and plasmid DNA purified by a miniprep procedure. If purification systems, such as the Qiagen kit, are used the plasmid preparations can also be used for DNA sequencing after screening. The plasmids from the minipreps are screened to ensure that the expected amplified product size has been cloned, by cutting out the insert with restriction enzymes and running the digests on a 1.5% agarose gel. Plasmids containing the correct size insert are then sequenced by conventional dideoxynucleotide sequencing, either manually or with an automated sequencer. The insert is sequenced from both directions (opposite strands).
Phylogenetic analysis The nitrogenase DNA sequences from both strands are compared and ambiguous positions resolved by re-examining the sequence data or resequencing, if necessary. The vector and PCR primer sequences are deleted from the consensus sequence from both strands. The sequence is then ready for analysis and submission to GenBank. The DNA or the deduced amino acid sequences can be used for phylogenetic analysis (Figure 14.2). However, the DNA sequence analysis is biased by the GC content of the third position of the codons, and eliminating the third position in the analysis results in a tree that is similar to that obtained with the deduced amino acids (Zehr el al., 1997). Given that the DNA sequences are a bit harder to align than the deduced amino acid sequences, the amino acid sequences are generally preferable for phylogenetic analysis. The DNA sequences can be translated on the World Wide Web (WWW) at the ExPasy server (http://www.expasy.ch/). The amino acid sequences (which are just over 100 residues long) can be manually aligned using alignment software (such as the Genetics Computer Group, GCG, software), or aligned computationally (CLUSTAL, or PILEUP in the GCG package). Computer alignments should be inspected carefully and adjusted manually. A number of phylogenetic packages are available, including distance methods in the PHYLIP package, parsimony in the PHYLIP package, PAUP or GCG and maximum likelihood (see the PUZZLE program http://members.tripod.de/korbi/puzzle/). GCG is a good program for the UNIX platform, PAUP for the Macintosh, and TREECON for the PC. However, there are now a variety of choices, most of which can be found on the WWW. The results of a number of the analysis packages result in the same basic features of the phylogenetic trees. Bootstrapping is important for interpreting the trees, since often some aspects of the tree topology are not well-supported by the analysis. The phylogeny of nifH is characterized by at least four major clusters. These clusters include conventional and vanadium nifH genes from
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proteobacteria and cyanobacteria in Cluster I, nifH sequences from second alternative nifH and nifH from Archaea in cluster II, and nitrogenases from anaerobes in Cluster III. Cluster IV includes very divergent n/f-like genes that have been found in Archaea. In general these groups are supported by high bootstrap values and sequences from the environment can be identified as being from these groups, or even specific groups within Cluster I. Sometimes, however, clusters of sequences fall in positions distant from the other groups such that placement within these larger clades is difficult. For example, the sequence group from aquatic environments described by Zani (Zani et al., 2000) group with Cluster I with a high bootstrap value (Figure 14.2), but the length of branches suggest some uncertainty as to their phylogenefic association. Such sequences will probably become clear as more sequences become available from known cultivated microorganisms. Such uncertainty in positioning can occur within individual clusters as well.
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NITROGENASE GENE EXPRESSION
Principle Genes are transcribed into messenger RNA molecules (mRNA) that are translated into protein. Transcription is the first step in synthesizing enzymes and structural proteins. The presence and abundance of specific mRNA molecules in the cell is controlled by the relative rates of transcription and degradation, and changes in the abundance of mRNA can reflect transcription rates of individual genes. There are a number of methods for assaying the presence of mRNA, from conventional Northern blotting, to S1 nuclease techniques and RT-PCR. RT-PCR has the advantage that it is a very sensitive technique, as well as being very specific. RT-PCR has been used for many different types of applications. For studies of expression by mixed populations, such as assemblages of nitrogen-fixing microorganisms in the environment, RT-PCR also has the advantage that mixed amplification products (derived from different microorganisms in the sample) can be characterized by subsequent cloning and DNA sequencing, or DNA probe hybridizations. Overall, the approach is as outlined for assaying nif genes, except that the resulting data indicate that organisms were synthesizing the nitrogenase enzyme, rather than simply having the nil gene in the genome.
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Equipment The equipment needed is as listed above for PCR.
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Reagents Reagents needed are diethyl pyrocarbonate (DEPC), bleach, 3% hydrogen peroxide, RNase Zap (Ambion), and a RT-PCR kit (Promega).
Methods Procedures for maintaining an RNase-free environment must be carefully followed. All glassware should be bleached and rinsed with RNase-free water. Soak all plasticware in 3% hydrogen peroxide for 12 h and rinse with RNase-free water. Pipette tip boxes and microcentrifuge tube racks can be treated with RNase Zap. All glassware, tips and solution containers should be autoclaved for a full hour then dried overnight in a drying oven at 37°C. Treat all solutions (except those containing Tris) with DEPC (0.1%) shaking gently and incubating for 2 h at 37 ° C. Autoclave solutions for 1 h to destroy the DEPC. Make Tris solutions with DEPC-treated water. All electrophoresis tanks should be washed with 0.5%SDS, rinsed with RNase-free water, rinsed with ethanol and dried. Store RNA in water at -20 ° C or -70 ° C. There are numerous methods and kits for processing RNA samples. The method described here was successfully used in a study of planktonic nitrogen-fixers (Zani et al., 2000). Total RNA can be extracted from filters using the RNeasy mini kit (Qiagen, Valencia, California). In this method, the extracted RNA is purified with a mini spin column (Qiagen, Valencia, California) according to the manufacturer's protocol and then resuspended in 50 gl H20. DNA in the samples can then be removed by digestion with DNase for 30 min at 37°C. The DNase enzyme is then removed from the sample using the RNeasy mini kit (Qiagen, Valencia, California) protocol. This protocol is based on the reverse transcription reaction using AMV reverse trancriptase (Promega, Madison, Wisconsin). The reverse transcription (RT) reactions are performed in 28 ~tl DEPC-treated H20, 10 gl AMV 5x buffer, 1 gl dNTP mix (10 mM each dNTP) and 1 pmol of primer nifH3. Prior to the RT reaction, the tubes are exposed to UV light (254 nm) for 20 min to prevent contamination. After irradiation, AMV reverse transcriptase (1 gl) and 1 ~1 of the DNase-treated RNA sample is added. Reactions are incubated at 42 oC for 30 min, and 1 gl of the resulting cDNA is added to a PCR mix (4 mm MgCL, 10x reaction buffer, 10 mM dNTPs, including 100 pmol of nifH3 and nifH4 primers and 2.5 U Taq polymerase) for the first round of nested PCR. The polymerase chain reaction is performed for 25-30 cycles of denaturation at 95 °C (1 min), annealing at 55°C (1 min), and extension at 72°C (1 min). The second nested PCR is carried out with 1 gl of the first round product using the same conditions as above, except by replacing primers nifH3 and nifH4 with primers nifH1 and nifH2. Samples can be tested for the presence of contaminating DNA in the RNA samples by performing nested PCR without the initial reverse transcription step (Figure 14.3). The samples can also be RNased to confirm that the amplification resulted from the RNA in the sample, as a second 282
test for contamination (Figure 14.3). These controls should be performed with each set of RNA extractions and RT-PCR reactions.
PROBLEMS, L I M I T A T I O N S A N D CAVEATS There can be problems with this approach in samples from complex matrices, or in samples that have few nitrogen-fixing microorganisms. The most common problem with this approach is obtaining a sample that does not inhibit the PCR. The nested PCR approach has greatly reduced this problem in samples that either have very low numbers of nitrogenfixing microorganisms or that have a high level of inhibitory compounds. Often, beginning experimenters have difficulty with multiple amplification products with these primers. We have found that high purity of the primers is essential for this approach. However, the nested PCR has greatly aided in the reduction of non-specific products as well. It is often the case, particularly with one-step nifH PCR, that a non-nif non-specific product of approximately the same size as nifH is amplified. This can be determined by cloning and sequencing some of the amplification products. Controls are crucial for PCR as well as RT-PCR. A positive control should be included and run in the sample matrix if competitive PCR is not being used to determine the extent of inhibition by the sample matrix. Negative controls are imperative for interpretation of the results. The use of cloned PCR products for positive controls creates a danger of crosscontamination. The procedures outlined here should be performed to alleviate contamination problems, but the possibility of contamination should be considered throughout the process, from sample handling to data analysis. A primary concern in the application of RT-PCR is ensuring that the amplified products are derived from reverse transcription and amplification of messenger RNA, and not from DNA. Appropriate controls help to ensure that the observed amplification is from the RNA. c o
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The PCR approach has been used in many environmental applications to study a number of different genes. Nitrogenase has received increasing attention and is a particularly good candidate for PCR approaches in terrestrial (Widmer et al., 1999; Ueda et al., 1995; Picard et al., 1992; Ohkuma et al., 1996) and marine environments (Bagwell et al., 1998; Zehr and McReynolds, 1989; Ben-Porath et al., 1993; Kirshtein et al., 1991; Zehr et al., 1995; 1998; Braun et al., 1999). PCR provides the basis for a number of approaches for quantifying and identifying nitrogen-fixing microorganisms and determining the diversity of microorganisms present (Zehr et al., 1995). Results of studies of plankton (Zehr et al., 1998) microbial mats (Zehr et al., 1995) and seagrass sediments (Kirshtein et al., 1991; 283
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Bagwell et al., 1998) have shown that there are diverse nitrogen-fixing microorganisms present in a variety of marine environments. Furthermore, different sets of nif genes are found in different habitats. Most striking is the presence of nif sequences from anaerobe microorganisms which are found in sediments (Bagwell et al., 1998), mats (Zehr et al., 1995) and associated with invertebrates (Braun et al., 1999), but are not found in picoplankton (Zehr et al., 1998) from the open ocean. The pattern of distribution of microorganisms indicates that there is selection for specific groups of diazotrophs in different environments, although it remains to be seen how many of these microorganisms actually express the nitrogen fixation apparatus. Results of a study of expression of niflq in natural assemblages in a freshwater lake indicate that many of the phylotypes detected by PCR may also be expressing nitrogenase genes (Zani et al., 2O0O).
FUTURE DIRECTIONS AND ALTERNATIVE APPROACHES Techniques are being developed that make it possible to amplify and detect genes. Some of these techniques include in situ techniques which make it possible to visualize cells microscopically that contain specific genes or messenger RNA molecules (Hodson et al., 1995). Amplification assays involving fluorescent assays such as molecular beacons or Taqman assays also promise new levels of specificity of detection and quantification. DNA probe hybridization techniques for characterizing nff amplification products have not yet been exploited to their full potential (Ben-Porath et al., 1993). Rapid methods for characterizing mixed assemblages of amplified DNA and RNA sequences, such as DGGE and T-RFLP, will greatly facilitate studies of the temporal dynamics and spatial distribution of nitrogen-fixing phylotypes (Bagwell et al., 1998; Ohkuma et al., 1999). Molecular hybridization strategies undoubtedly will ultimately be used on DNA chips to provide molecular technologies for the study of nitrogen fixation in the environment.
References Bagwell, C. E., Piceno, Y. M., Ashburne-Lucas, A. and Lovell, C. R. (1998). Physiological diversity of the rhizosphere diazotroph assemblages of selected salt marsh grasses. Appl. Environ. Microbiol. 64, 4276-4282. Ben-Porath, J., Carpenter, E. J. and Zehr, J. P. (1993). Genotypic relationships in Trichodesmium (cyanophyceae) based on nifH sequence comparisons. I. Phycol. 29, 806-810. Braun, S., Proctor, L., Zani, S., Mellon, M. T. and Zehr, J. P. (1999). Molecular evidence for zooplankton-associated nitrogen-fixing anaerobes based on amplification of the nifH gene. FEMS Microbiol. Ecol. 28, 273-279. Burlage, R. S. (1998). Molecular techniques. In: Techniques in Microbial Ecology (R. S. Burlage, R. Atlas, D. Stahl, G. Geesey, and G. Sayler, Eds), pp. 289-336. Oxford University Press, New York. 284
Howard, J. B. and Rees, D. C. 1996. Structural basis of biological nitrogen fixation. Chem. Rev. 96, 2965-2982. Hodson, R. E., Dustman, W. A., Garg, R. M. and Moran, M. A. (1995). In situ PCR for visualization of microscale distribution of specific genes and gene products in prokaryotic communities. Appl. Environ. Microbiol. 61, 4074-4082. Kirshtein, J. D., Paerl, H. W. and Zehr, J. R (1991). Amplification, cloning, and sequencing of a nifH segment from aquatic microorganisms and natural communities. Appl. Environ. Microbiol. 57, 2645-2650. Larrick, J. W. (1997). The PCR Technique: Quantitative PCR. Eaton Publishing. Ohkuma, M., Noda, S., Usami, R., Horikoshi, K. and Kudo, T. (1996). Diversity of nitrogen fixation genes in the symbiotic intestinal microflora of the termite Reticulitermes speratus. Appl. Environ. Microbiol. 62, 2747-2752. Ohkuma, M., Noda, S. and Kudo, T. (1999). Phylogenetic diversity of nitrogen fixation genes in the symbiotic microbial community in the gut of diverse termites. Appl. Enviroll, Microbiol. 65, 4926-4934. Picard, C., Parsonnet, E., Paget, E., Nesme, X. and Simonet, P. (1992). Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction. Appl. Environ. Microbiol. 58, 2717-2722. Ueda, T., Suga, Y., Yahiro, N. and Matsuguchi, T. (1995). Remarkable N:fixing bacterial diversity detected in rice roots by molecular evolutionary analysis of izi/CHgene sequences. J. Bact. 177, 1414 1417. Widmer, E, Shaffer, B. T., Porteous, L. A. and Seidler, R. J. (1999). Analysis of ~lifH gene pool complexity in soil and litter at a Douglas fir forest site in the Oregon Cascade mountain range. Appl. Environ. Microbiol. 65, 374-380. Zani, S., Mellon, M. T., Collier, ]. and Zehr, J. P. (2000). Application of a nested reverse transcriptase polymerase chain reaction assav for the detection of iffflt expression in Lake George, New York. Appl. Etlviron. Micn~biol. 66, 3119 3124. Zehr, J. P. and McReynolds, L. A. (1989). Use of degenerate oligonucleotides for amplification of the nifH gene from the marine cyanobacterium Trichodesmium spp. Appl. Environ. Microbiol. 55, 2522-2526. Zehr, J. P., Mellon, M., Braun, S., Litaker, W., Steppe, T. and Paerl, H. W. (1995). Diversity of heterotrophic nitrogen fixation genes in a marine cyanobacterial mat. ANd. Environ. Microbiol. 61, 2527-2532. Zehr, J. P., Melk)n, M. T. and Hiorns, W. D. (1997). Phylogeny of cyanobacterial n!fiH genes: evolutionary implications and potential applications to natural assemblages. Microbiology 143, 1443-1450. Zehr, J. P., Mellon, M. T. and Zani, S. (1998). New nitrogen fixing microorganisms detected in oligotrophic oceans by the amplification of nitrogenase (nifH) genes. Appl. EJlviroH. Microbiol. 64, 3444-3450. Zimmermann, K. and Mannhalter, ]. W. (1996). Technical aspects of quantitative competitive PCR. Bio Teclmiques 21, 268-279.
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List of suppliers PHYLIP software at http://evolution.genetics. washington.edu/phylip.html
Bio Rad Laboratories 2000 Alfred Nobel Drive Hercules, CA 94547, USA 800-4-BIORAD
Promega Fisher Scientific
2800 Woods Hollow Road Madison, W153711, USA 800-356-9526
4500 Turnberry Drive Hanover Park, IL 60103, USA 630-259-1200
PUZZLE software at http://members.tripod.de/korbi/ puzzle/
Genetics Computer Group 575 Science Drive Madison, W153711, USA 608-231-5200
Qiagen Inc. 28159 Avenue Stanford Valencia CA 91355, USA 800-426-8157
New England Biolabs, Inc. 32 Tozer Road Beverly, MA 01915, USA 800-632-5227
Novagen 601 Science Drive Madison, WI53711-1084, USA 608-238-6110
PE Applied Biosystems 850 Lincoln Center Drive Foster City, CA 94404, USA 800-345-5224
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TREECON software at http://bioc-www.uia.ac.be/u / yvdp / treeconw.html
15 Protistan Herbivory and Bacterivory David A C a r o n Department of Biological Sciences, 3616 Trousdale Parkway, AHF 30 I, University of Southern California, Los Angeles, California 90089-037 I, USA
CONTENTS
Introduction General theory and approaches Protistan herbivory measured using the dilutiontechnique Protistan bacterivory measured using FLB Conclusions and recommendations
INTRODUCTION Marine protists comprise a highly diverse collection of species that play numerous ecological roles. These single-celled, eukaryotic organisms vary in size from ~-2 lam to more than 1 mm, although some colonial species also form conspicuous macroscopic structures. Traditionally, ecologists have divided protists into photosynthetic forms (the microalgae or phytoplankton) and heterotrophic forms (the protozoa). In this scenario, chloroplast-containing protists obtain their nutrition via the uptake of inorganic compounds and photosynthetic carbon fixation, while apochlorotic species obtain nutrition via the engulfment (phagocytosis) of prey and possibly by direct uptake of dissolved organic compounds from the surrounding medium. Protists capable of phagocytosis form a major component of the consumer assemblages in marine ecosystems. These protists constitute a significant portion of the standing stock of microbial biomass in pelagic ecosystems (Stoecker et al., 1994; Caron et al., 1995; Stoecker el al., 1996; Dennett et al., 1999), and they are responsible for the consumption of large amounts of primary and bacterial production (Sanders el al., 1992; Sherr and Sherr, 1994). Thus, they occupy a pivotal role as consumers in aquatic food webs (Azam et al., 1983; Fenchel, 1987; Caron and Swanberg, 1990; Caron, 1997), and form an important source of food for many planktonic metazoa (Stoecker and Capuzzo, 1990; Sanders and Wickham, 1993). Phagotrophic protists also contribute significantly to the remineralization METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0-12-521530 4
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of macro- and micronutrients in aquatic ecosystems (Goldman and Caron, 1985; Barbeau et al., 1996). While the heterotrophic activities of protists in aquatic ecosystems are unquestionably important, accurate quantification of the rates of herbivory and bacterivory conducted by these assemblages has been difficult. This difficulty is partly due to the diversity of size and trophic activity of protists. The smallest phagotrophic protists are capable of consuming only bacteria and cyanobacteria, while the largest (some foraminifera and actinopods) can consume prey larger than 1 mm. Trying to devise an experimental design that will apply over these tremendous ranges of size and nutritional modes is problematic. Another important factor complicating the accurate determination of heterotrophic activity by aquatic protists is that many species are capable of mixed nutrition. Numerous recent publications and several reviews have indicated that protistan nutrition is not nearly as straightforward as simple, alternative behaviors of photosynthesis and heterotrophy. Many of these species possess the ability to combine these two modes in a single individual. Numerous species of chloroplast-bearing flagellates possess the ability to ingest and digest particulate prey in addition to conducting photosynthesis, while some ciliates that prey on algae can retain the chloroplasts of their prey in a functional state for a limited period of time. These behaviors have resulted in situations where a significant fraction of the bacterivory in some aquatic ecosystems has been attributed to phytoplankton (Berninger et al., 1992), or to instances where protozoa have been shown to contribute significantly to primary production (Crawford, 1989). In addition, mutualistic endosymbioses involving photosynthetic microorganisms held within the cytoplasm of heterotrophic protists are abundant in some marine ecosystems. Taken together, it is becoming increasingly clear that these behaviors and symbiotic relationships are much more common than previously believed, and that they add significantly to the structure of microbial food webs (Sanders, 1991; Jones, 1994; Caron, 2000). The existence of these complex nutritional modes among protistan species confounds our often simplistic view of protistan nutrition, and complicates our ability to neatly and accurately measure heterotrophic processes by these organisms. For this reason, the older terminology that categorizes protists according to autotrophic or heterotrophic nutrition ('algae', 'protozoa') is losing popular appeal among ecologists dealing with energy flow through microbial food webs. A more appropriate term 'phagotrophic protists' is gaining popularity with reference to species capable of ingesting prey (regardless of the presence or absence of chloroplasts). Nevertheless, historical precedence for the older terminology is strong, and one still finds these terms in common use (Andersen, 1992; Hausmann and H/ilsmann, 1996). One final consideration when examining heterotrophy by protistan assemblages stems from the need, for most methods, to make relatively long observations of captive populations. However, merely placing microorganisms in bottles for making physiological measurements may result in aberrant behavior, death or increased growth (depending on the 290
species). For example, the enhancement of bacterial activities during containment, the so-called 'bottle effect', has been known for quite some time (Zobell, 1943). The effects of containment must be weighed when evaluating the results of some experimental approaches. The issues described above have resulted in a variety of experimental approaches for estimating rates of protistan herbivory and bacterivory. It is important to stress that each of these approaches has its purpose, advantages and disadvantages. Therefore, it is necessary to consider the specific characteristics of each method, and the attributes of the ecosystem to be investigated, in choosing the most appropriate approach to reach the experimental goals. Two experimental protocols commonly used to examine bacterivory and herbivory by phagotrophic protists are described in this chapter. Preceding the description of each method is a short discussion of the theory, advantages and disadvantages of experimental approaches for measuring protistan food web interactions. A number of variations on these methods exist as well as alternative approaches. Reference to alternative protocols and variations on these two methods are provided as space permits. For a more complete discussion of other methodologies, the reader is referred to some recent reviews (Caron et al., 1993; Sherr and Sherr, 1994; 1997).
G EN E R A L TI-I EO RY A N D A P P R O A C H ES Experimental approaches for measuring protistan trophic activity (herbivory and bacterivory) can be characterized according to the taxonomic level at which they are directed, and the general approach taken to assess the activity of the assemblage. That is, such measurements may be directed toward a particular species within an assemblage ('taxonspecific'; e.g. feeding rates of a particular ciliate species on bacteria), or toward the entire assemblage ('community-level'; e.g. total rate of bacterivory taking place in a water sample). This decision (and the nature of the species involved) will affect the choice of the experimental protocol and specific aspects of its application (e.g. incubation time). In addition, experimental designs can be classified into one of two general approaches; (1) perturbation of trophic relationships and monitoring consequent changes in prey abundance/biomass, or (2) tracer experiments that attempt to follow the uptake of prey into consumers or their removal from the water.
P e r t u r b a t i o n e x p e r i m e n t s : pros and cons There is a fairly long history of measuring rates of protistan herbivory and bacterivory by perturbing the grazing activities of these minute predators which have yielded a number of specific approaches. A fundamental assumption is that prey populations are growing and dying (being consumed) in natural microbial assemblages. At their core, all of these
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methods share the common theme of somehow uncoupling prey growth from predation, and using the resulting changes in prey abundances to estimate the impact of their consumers. The most basic of these approaches involves size fractionation of water samples, and the subsequent measurement of changes in prey abundance or biomass in the filtrate and in the unfractionated water sample. This method assumes that predators are significantly larger than their prey and therefore can be removed efficiently by filtration. In this scenario, changes in the prey population in the filtrate (predators removed) relative to changes in the unfractionated water sample (predators present) are indicative of the amount of prey removed by consumers. Fractionation experiments offer the advantages that they are relatively straightforward to conduct, and that they rely on natural prey/predator interactions (i.e. no artificial prey or tracers are employed; see next section). This approach is rarely applicable for examining the trophic activity of individual species because it is usually not possible to remove individual species by filtration. In most cases this approach provides a 'community level' estimate of grazing activity. Problems can occur, however, when the sizes of protistan predators and prey overlap. This problem is particularly acute for rates of herbivory because primary producers and phagotrophic protists both span wide, overlapping size ranges. In some cases protistan herbivores are smaller than their phytoplankton prey. For example, the longest dimensions of some heterotrophic dinoflagellates are significantly less than those of their diatom prey (Jacobson, 1999). Size fractionation has been used most effectively for examining bacterivory (or cyanobacterivory) because the latter prey assemblages can be separated more efficiently from their predators (Wright et al., 1987; Caron et al., 1989; Weisse and Scheffel-M6ser, 1991). Another potential problem of size fractionation approaches is that prey growth may be dependent on substrates or nutrients released by consumers. Removal of the latter by filtration may adversely affect the growth rates of prey (relative to their growth rates in unfiltered water). Interruption of this 'feedback loop' can result in underestimation of the growth of prey when predators are removed, and is a potentially serious problem for this method (Sherr et al., 1988). Chemical perturbation of microbial activities (rather than physical separation) also has been employed to examine bacterivory by phagotrophic protists. Like size fractionation, this approach allows experimentation with natural predator/prey populations (i.e. no artificial prey). A common approach is the use of metabolic inhibitors that selectively inhibit eukaryotic cell function (e.g. colchicine, cycloheximide) and thus stop the grazing activities of protists. A comparison of the increases in bacterial (prey) abundances in the presence of these inhibitors (relative to changes in water samples without the inhibitors, where grazing continues to take place) provides an indication of the grazing impact of the eukaryotes (Campbell and Carpenter, 1986; Sherr et al., 1986; Caron et al., 1989). Bacterial inhibitors also have been employed occasionally. In the latter situations, decreases in prey abundance (relative to controls without 292
inhibitors) are monitored. Metabolic inhibitors of photosynthesis (e.g. DCMU) have not been used routinely to examine protistan herbivory. Caveats for the use of metabolic inhibitors include interruption of a feedback loop as mentioned above by interrupting grazer metabolism, the potential for heterotrophic bacteria to use some eukaryotic inhibitors as growth substrates, and the potential for bacterial inhibitors to affect eukaryotic growth processes (i.e. non-specific inhibition). For these reasons, metabolic inhibitors are no longer commonly used in studies of microbial predation. One type of perturbation experiment that is increasingly employed to examine microbial (presumably protistan) herbivory in water samples is the dilution technique (Landry and Hassett, 1982; Landry, 1993; Landry et al., 1995b). This method relies on the dilution of protistan grazers with filtered water in a series of experimental bottles in order to progressively reduce grazing pressure on the phytoplankton. The method is described in more detail below.
T r a c e r experiments: pros and cons An alternative approach to perturbation experiments relies on the use of tracers to follow the ingestion (or removal) of microbial prey. This approach is designed to minimize some of the inherent difficulties associated with perturbation experiments, particularly the interruption of feedback loops within the microbial communities. Tracer experiments often use specific compounds (typically radioactive or fluorescent entities) to label natural prey populations or assemblages, thereby allowing their biomass to be monitored and quantified as it moves through food webs. Alternatively, pre-labeled prey (cultured prey or even artificial prey) may be added to a natural water sample with the assumption that these labeled populations will be consumed by microbial predators at rates equivalent to similarly-sized, naturally-occurring prey. Fluorescent prey 'mimics' that have been used include fluorescently labeled algae (Rublee and Gallegos, 1989) for measuring rates of herbivory, or fluorescently labeled bacteria (FLB), paint particles or plastic beads for measuring rates of bacterivory (McManus and Fuhrman, 1986; Pace and Bailiff, 1987; Sherr et al., 1987; McManus and Okubo, 1991). The use of FLB to examine bacteriw)ry has become a common tool in microbial ecology, and this method is described in detail below. The use of radiolabeled prey has unique advantages and disadvantages for measuring rates of protistan grazing relative to other approaches. Isotopes are available that can effectively label either photosynthetic or bacterial assemblages (e.g. H"CO~ for labeling phytoplankton, 'H-thymidine (TdR) for bacteria). Radiolabeling provides very high sensitivity, enabling the possibility of analyzing the trophic activity of the entire consumer assemblage, or individual protistan cells (Taylor and Lean, 1981; Lessard and Swift, 1985). Thus, these approaches can be modified to gather taxon-specific information about consumers, as well as community-level grazing impact of the consumers (although not always
293
e-
l.
0
0
in the same experimental set-up). In addition, these experiments typically require minimal manipulation of the microbial assemblages that are being examined (compared to filtration, dilution or the use of inhibitors). Prelabeled prey can be added to untreated water samples, and direct additions of some radiolabels to water samples are possible for situations where the label is specifically taken up by prey (e.g. the use of TdR to label bacteria). Finally, radiolabeling methods can be modified to yield information on metabolic processes of the consumers (Taylor and Sullivan, 1984; Zubkov and Sleigh, 1995). While radiolabeling methods are potentially powerful approaches for examining protistan grazing activities, they also have some distinct drawbacks. A practical consideration in radiolabeling experiments is following the movement of the label from prey to consumers. This goal is generally accomplished by filtration of the assemblage onto a filter that will retain the consumers (and ingested prey) but allow uningested prey to pass. Analysis of the radioactivity retained on the filter (normalized to the specific activity of the prey) indicates the amount of prey biomass consumed. Unfortunately, clean separation of prey and protistan consumers is problematic as noted above, and thus considerable contamination of the 'consumer fraction' with uningested prey can occur. These errors result in the overestimation of trophic activity by phagotrophic protists. Micropipetting individual protistan cells for analysis overcomes this problem, but it is a highly laborious and exacting procedure (Lessard and Swift, 1985). An important theoretical consideration for radiolabeling experiments is the potential artefact of label release a n d / o r recycling during the experiment. Protists can release or remineralize a large proportion of their prey biomass during relatively short incubations (e.g. approx. 1 h). Therefore, the timing of sample collection and processing is critical in radiolabeling experiments in order to avoid considerable underestimations in protistan grazing rates (Caron et al., 1993). Finally, isotope use, containment and disposal present increasingly difficult challenges in conducting field experiments, and thus the use of radiolabeling approaches for examining protistan bacterivory and herbivory are better suited for controlled conditions in the laboratory.
~t~tt~
P R O T I S T A N HERBIVORY MEASURED U S I N G THE DILUTION TECHNIQUE
Principle The dilution technique was first proposed by Landry and Hassett (1982). Since that time it has become increasingly popular for estimating the rate of herbivory by microzooplankton (i.e. zooplankton <200 ~m, a size class typically dominated by protists). This method has been applied successfully in a wide range of freshwater and marine ecosystems (estuarine to oceanic), including tropical, temperate and polar environments. Subsequent refinement of the technique from the original description
294
(Landry et al., 1995b) has resulted in changes that attempt to alleviate the possibility of grazer-dependent growth of the phytoplankton within the dilution series. A unique result of the dilution method, as presently employed, is that it allows the simultaneous measurement of community-level growth rate of the phytoplankton as well as community-level mortality rate (herbivory). The experimental protocol entails the establishment of a series of water samples that are diluted to varying degrees with filtered (0.2 !am) water. In practice the dilution series typically has between five and seven treatments (in triplicate) varying from 100% to 20% unfiltered water. The entire dilution series receives inorganic nutrients and trace metals to promote equivalent gross growth rates of the phytoplankton assemblage in all experimental bottles. A replicate set of bottles containing 100% unfiltered water is also prepared but not enriched with nutrients. The latter treatment serves as a measure of natural (i.e. unenriched) phytoplankton growth and mortality rates in the water (Figure 15.1). All bottles are incubated, and phytoplankton growth and mortality are determined from changes in phytoplankton biomass in the various treatments. It is assumed in the dilution method, as it is applied routinely, that phytoplankton growth in the bottles is independent of the presence or absence of grazers, and that clearance rates of the herbivorous phagotrophic protists are unaffected by dilution. The mortality rate (m) is indicative of the grazing losses (herbivory) in each bottle assuming that herbivory dominates mortality of the phytoplankton assemblage. Grazing impact of the herbivores in each treatment is dependent on their absolute abundances, which in turn is dependent on the degree of dilution with filtered water (D~). Thus, net phytoplankton growth rate (ki) in each enriched dilution bottle is a consequence of the growth rate of the phytoplankton with enrichment (]_l,,, assumed to be constant in all enriched bottles), the mortality rate (m) and the relative abundance of consumers. ki = la,,- (m x D)
(15.1)
Net phytoplankton growth rates are calculated for each bottle of the enriched dilution series and for the unenriched bottles. The simplest means of accomplishing this task is to assay chlorophyll a fluorometrically at the beginning and end of the incubation period in each of the bottles, and calculate apparent phytoplankton growth rates in each bottle according to the equation k, = (ln ChlEf I - In C h l , ~ l ) / ( i n c u b a t i o n
time)
(15.2)
where Chllo ~ and Chlff~ are the chlorophyll concentrations at the beginning (time = 0) and end (time = final) of the incubation, respectively. This calculation assumes exponential growth of the phytoplankton. Other methods of analyzing phytoplankton growth rates can be used (see Caveats, precautions and alternatives). Mortality rate is obtained from the slope of the regression of phytoplankton growth in each bottle of the enriched dilution series plotted 295
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100% Unfiltered water (+ nutrients) 80% Unfiltered water 20% Filtrate (+ nutrients)
~
~
60% Unfiltered water 40% Filtrate (+ nutrients)
~
40°/°water Unfiltered
Following incubation (usually for 24 h). Resample all bottles for chlorophyll. Calculate net phytoplankton growth rate (kl) in each treatment from changes in chlorophyll. Express k graphically as a function of the fraction unfiltered water.
60% Filtrate (+ nutrients) O ° o Uwn l etr
80% Filtrate (+ nutrients)
@
~
100% Unfiltered water (no nutrients)
Followin~ incubation (usually for 24 h). Resample all bottles for chlorophyll. Calculate net phytoplankton growth rate (ko) from changes in chlorophyll during incubation.
Figure 15.1. Overview of a typical layout for the dilution method (Landry et al., 1995b) for measuring phytoplankton growth rate and mortality rate (herbivory). A series of mixtures of unfiltered and filtered water are prepared and incubated, Grazing mortality increases with the proportion of unfiltered water.
296
against the fraction of unfiltered water in each bottle (Figure 15.2). Changes in phytoplankton biomass in the unenriched, undiluted treatment provide an estimate of the natural (i.e. unenriched) net phytoplankton growth rate (k,,). The instantaneous phytoplankton growth rate (p,,) in the water sample (i.e. in the absence of mortality and without nutrient enrichment) can be obtained by adding the mortality losses to net growth rate. That is, (15.3)
t~L, = k , + m
This terminology is consistent with recent applications of the method (Landry et al., 1995a; 1995b; 1998; Caron and Dennett, 1999). 1.2
Slope = Mortality rate (m; units of d4)
~ 0.8 f..-
o c~ 0.6
¢.. 0
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~_
0.4
0 13-
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Net phytoplankton growth rate (ko; units of d 1) J
0
I
I
I
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0.4
0.6
0.8
Relative grazing pressure
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Figure 15.2. Example of the results obtained using the dilution method to measure phytoplankton growth rate and mortality rate. Pertinent features shown are the net phytoplankton growth rate in the unenriched (k) and enriched (k.) treatments, and the mortality rate (m, slope of the regression).
L
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O I. D.
Equipment and reagents • Collection, preparation and incubation apparatus: 50 I carboy with tubing ports in cap, 20 I carboy with spigot; silicone tubing; black plastic; graduated cylinders.
297
• An acid-washed, pre-rinsed 0.2 pm cartridge filter (Gelman ® filters work well), and 200 IJm nylon screening held in an in-line filter cartridge. • Dilution bottles; minium of 21, I-2 I, clear, polycarbonate bottles.This number of bottles assumes five enriched dilution treatments (e.g. 20, 40, 60, 80 and 100% unfiltered water), a set of unenriched 100% bottles, and a set of controls (diluent only), all in triplicate. • Coolers for holding and transferring dilution bottles. • Sufficient darkened sample bottles for collection of chlorophyll samples (two bottles for each dilution bottle). • Chlorophyll analyses: multiple filtration rigs (25 mm base, large volume tower) and filtration manifold; vacuum pump; glass fiber filters (GF/F Gelman®); forceps, acetone; repipettor for acetone; dilute HCI; eye dropper; I 0 ml glass tubes with caps (for extractions); aluminum foil; fiuorometer. • Macronutrient and trace metal stock solution: Nitrogen (NH4CI), phosphorus (Na2HPO/7H~O), iron (FeCI) and manganese (MnSO,); micropipettor. • Equipment for in situ incubation of dilution bottles (e.g. clear plexiglass platform; bottle fasteners, etc.). Alternatively, an outside incubator at ambient water temperature is often employed (at sea this can be kept at ambient temperature by continuously pumping seawater through it), equipped with clear sides and gray window screening to reduce light to ambient levels (or colored, plastic sheets to mimic light quality and quantity). • Computational facilities for analyzing data and plotting regressions. • Samples are usually collected and preserved for microscopy, flow cytometry or HP/C in addition to chlorophyll. Additional bottles (and appropriate preservatives) should be added as necessary.
Experimental setup All materials used to collect, filter, transfer or hold water samples should be acid-washed in 5-10% HC1. All transfers involving unfiltered water should be performed with an absolute minimum of bubbling. Bubbles can cause significant damage to delicate plankton. Silicone tubing should be employed to avoid the toxic effects of some plastics. Care also should be taken not to expose water samples to bright light (containers can be covered with black plastic), or temperatures above or below ambient temperature when preparing the dilution series. Transfer the incubation bottles to and from the incubation site in a cooler. 1. Fill the 50 1 carboy with water from the experimental site/depth and remove the carboy to a suitable location to prepare the dilution series. Prepare the diluent by gravity filtration using silicone tubing and the 0.2 l~m pore-size, acid-washed cartridge filter. Cartridge filters have excellent filtration capacity, and pre-filtration should be necessary only in highly turbid ecosystems. 2. Dispense the appropriate amount of diluent into each of the dilution bottles such that filling the remainder of the bottle to the neck with unfiltered water will produce the desired dilution. For example, a typical dilution series might entail bottles with 20, 40, 60, 80 and 100% unfiltered water (other dilution schemes can be used). For 2.0 1 incuba-
298
3.
4.
5.
6.
7.
tion bottles, this means 1600 ml of diluent and 400 ml of unfiltered water to obtain a treatment containing 20% unfiltered water. Fill all bottles to the appropriate level (usually the neck of the bottle) with unfiltered (actually, 200 pm filtered) water from the 50 1 carboy with minimal bubbling. The 'unfiltered' water is passed through 200 lJm screening in order to remove relatively rare mesozooplankton (mostly copepods). Therefore, herbivory in the <200 lJm filtrate is typically attributed to phagotrophic protists which dominate the nano(2-20 t~m) and micro- (20-200 lJm) zooplankton. Collect samples (a minimum of three, usually more) of the <200 pm water and from the <0.2 lJm diluent carboy for chlorophyll analysis (time = 0). A significant time saving can be realized if the volumes of the chlorophyll sampling bottles are accurately known when they are completely filled. This simple step obviates the need for measuring each sample every time. Therefore, it is advantageous to have several size bottles available and choose the volume appropriate for chlorophyll levels at the study site. Filter the entire contents of each chlorophyll bottle onto a glass fiber filter, place the filter into a glass tube, and extract by adding 7 ml of acetone, cover and place in the cold and dark. Convention dictates the use of 90% acetone, and storage in a refrigerator, although the use of 100% acetone and storage in a -20°C freezer works equally well (this avoids the need to dilute the acetone). Samples are read fluorometrically using standard techniques (Parsons et al., 1984) after 24h of extraction. Add an appropriate amount of the nutrient and trace metal stock solution(s) to each of the enriched bottles. The appropriate concentrations will vary with locale, but typical final concentrations of nutrients and trace metals for oceanic studies might be 10 I1M N, 1 I1M P, 1 nM Fe and 0.1 nM Mn. Tightly close all bottles, transfer them to the incubation site or incubator, and record time.
At the end of the incubation period (typically approx. 24 h), remove samples from all bottles for chlorophyll analysis (duplicate samples for all bottles) as described above.
Analyses Net phytoplankton growth rates in each of the enriched dilution bottles (k,, equation (15.2)) are plotted against the proportion of unfiltered water (i.e. relative grazing pressure) and analyzed with standard regression techniques (Landry and Hassett, 1982; Landry et al., 1995a). The negative slope arising from this regression is the mortality rate (in units of time '). The Y-intercept represents the instantaneous growth rates of the phytoplankton under nutrient-enriched conditions (p~,). The natural (unenriched) net phytoplankton growth rate (k,,) is indicated by the growth rate in the bottles with 100% unfiltered water but no nutrient amendation. This latter value and the mortality rate are used to calculate the instantaneous
299
O
growth rate of the phytoplankton in the water sample (i.e. ~,,; see equation (15.3)). A comparsion of phytoplankton growth rates in the undiluted bottles with and without nutrient enrichment provides additional information on the nutrient status of the phytoplankton assemblage. Significant upward shifts in net phytoplankton growth rate with the addition of nutrients and trace metals (i.e. ~n>>t~,,)is indicative of nutrient (or trace metal) limitation of the phytoplankton community in the water (Landry et al., 1998; Caron and Dennett, 1999).
Caveats, precautions and alternatives The dilution technique has now been used in a variety of marine (and freshwater) ecosystems, as noted above. A clear attraction of the method is that it is virtually the only method, at present, by which one can obtain estimates of phytoplankton growth rate and mortality rate simultaneously. In practice, it is probably most effective for small primary producers (e.g. the picophytoplankton; cells <2 ~m in size) because they and their grazers are more adequately sampled with relatively small sample sizes employed in the experimental protocol (_<21). For this reason it is used commonly in open ocean ecosystems. It is critical that the assumptions of the method be met in order to obtain valid data, and this is not an easy assessment to make. Erroneous conclusions (under- or overestimations of mortality rate) can be derived if phytoplankton growth rates vary along the dilution series (Landry, 1993), and obtaining useful data (i.e. significant slopes) at relatively low rates of mortality can be difficult. The basic method, as described here, utilizes measurements of chlorophyll as a proxy for phytoplankton biomass because it is a relatively straightforward analysis. However, this approach is not without its problems. Photoadaptation (i.e. changes in pigment content cell ') can occur during the incubation period, affecting growth rate calculations based on changes in chlorophyll (McManus, 1995). Therefore, incubating the dilution series in a light regime that does not match the natural light quality/quantity may yield phytoplankton growth rates that are inaccurate. This problem does not necessarily affect the mortality rate obtained unless photoadaptation is different within the dilution series. This latter situation seems possible only when phytoplankton are so dense that self-shading occurs at the lowest dilutions. A simple solution to this problem would be to incubate small sample volumes in containers that allow better light penetration (e.g. tissue culture flasks oriented toward the light source). It is also possible and advantageous to use supplemental or alternative means of assessing changes in phytoplankton abundance. Cell counts of phytoplankton in each bottle can be conducted, but the number of samples to be processed even in a single dilution experiment is rather prohibitive. Both HPLC and flow cytometry have been employed to follow changes in specific algal taxa (Burkill, 1987; Burkill et al., 1987; Latasa et al., 1997). These methods are more involved and require more
300
sophisticated equipment, but they also yield information on the growth and mortality of specific taxonomic components of the phytoplankton assemblage. A practical problem with the method is that it is very labor intensive, and a single experiment requires a minimum of three days to complete (setup on day 1, sampling and takedown on day 2, sample readings and data analysis on day 3). For this reason, and in order to minimize the effects of containment, incubation times should be as short as possible. However, they must be sufficiently long to allow measurable differences in net phytoplankton growth rates across the dilution series, in some ecosystems the latter requirement may necessitate quite long incubations (Caron et al., in press). Another consideration is that phagotrophic protistan assemblages may grow significantly during incubations within the dilution series, and potentially at different rates in response to food availability. These changes are difficult to examine in routine applications of the method. In the absence of information on the functional and population responses of the grazers in each experiment, it is generally assumed that individual clearance rates of the consumers are constant throughout the treatments of the dilution series. An experimental test of this assumption has indicated that, at least in some ecosystems, the assumption is valid (Landry et al., 1995b). However, previous attempts to use the method in eutrophic environments, and a more recent analysis of a large data set, indicate that feeding and growth of microzooplankton may vary within the dilution series (Gallegos, 1989; Dolan et al., submitted). Finally, the dilution technique has been used largely to examine microbial herbivory. In principle it can be applied to determine the growth and mortality rates of bacterial assemblages (Tremaine and Mills, 1987; Geider, 1989), but determining the substrate(s) and degree of organic enrichment to maintain bacterial growth within all dilutions is problematic. Bacterial growth rates can be highly responsive to the addition of labile organic compounds (relative to protistan growth and grazing rates), and maintaining growth without dramatically affecting bacterial abundance is a tricky matter.
PROTISTAN BACTERIVORY MEASURED U S I N G FLB
"O c-
Principle The use of fluorescently labeled prey to measure rates of microbial grazing has had many variations. All of these methods hinge on the same basic principle of using an easily identified (fluorescently labeled) prey or prey surrogate to trace the consumption of a microbial population or assemblage. The method described below entails the use of fluorescently labeled bacteria (FLB) produced from cultured bacteria. Microbial bacterivory is a difficult process to measure accurately in natural microbial assemblages. Tile choice of this method represents a good compro301
O
a.
raise between versatility and ease of performance on the one hand, and the efficacy of accurately assessing the ingestion rates of microbial bacterivotes on the other hand. There are two fundamental variations on the use of FEB that enable the measurement of either taxon-specific ingestion rates of bacteria or community-level mortality rates (Figure 15.3). Bacteria stained with the fluorochrome 5-(4,6-dichlorotriazin-2-yl) aminofluorescein (DTAF) were first produced by Sherr et al. (1987) as an alternative prey item to surrogate prey that had been used up to that time to examine the rate of ingestion of bacteria by phagotrophic protists (see Caveats, precautions and alternatives). In short, FEB are added to a water sample at a known concentration, and samples are removed periodically, preserved, and examined microscopically for the presence and number of ingested FEB (counterstaining the protists allows them to be visualized). Determination of the ratio of FLB to unstained, natural bacteria in the water sample, combined with information on the average rate of uptake of FEB by protists is then used to calculate the total rate of bacterial ingestion. This approach is ideal for determining the ingestion rates of protistan species that can be distinguished by light microscopy (Figure 15.4(a)). Because most bacterivorous species of protists cannot be identified by epifluorescence microscopy, however, ingestion rates of protistan 'morphotypes' are acquired in most cases (e.g. heterotrophic flagellates, phagotrophic phytoflagellates, oligotrichous ciliates). A 'community-level' ingestion rate can be obtained with considerable effort. A variation on this method uses the disappearance of FEB from the water as a community-level estimate of bacterivory. Studies have indicated that FLB are non-toxic and are readily metabolized by protists, and metabolism results in destruction of the fluorescent label (Sherr et al., 1987). Therefore, time series analysis of the rate of disappearance of FEB from a water sample indicates the rate of turnover of the bacterial assemblage in the water. Consumption is not attributed to any specific component(s) of the water sample in this approach, but total bacterivory is obtained (Figure 15.4(b)). The bacteria used to produce FEB can be obtained from cultured strains (singly or mixed), enrichments of bacteria from natural water samples (a 0.8 lJm filtrate is generally used to avoid the growth of protists), or natural bacteria concentrated from the environment. The choice is dictated somewhat by the breadth and goal of the study, and somewhat by practicality. Ideally, the size distribution of the FLB should be similar to the spectrum within the natural bacterial assemblage being studied, and this is best accomplished using concentrates of tile natural community to produce FLB. The process is laborious, requires large amounts of water and time, and suitable equipment for concentrating the bacteria (e.g. a tangential flow concentrator). In the end, each concentrate is applicable only to the water mass from which it came. Moreover, concentrating the bacteria results in some loss of cells, and ultimately the size spectrum of cells within the bacterial concentrate is not necessarily even a good reflection of the natural community from which it was taken. For this reason, cultured bacteria of various types often have been employed. 302
~R
FLB UPTAKE emove and preserve amples immediately and t specific time intervals.
~
Inoculate with FLB
Prepare samples for epifluorescence microscopy, counterstaining with DAPI.
Microscopically determine the number of ingested FLB per protist.
Nat::~mp~eate
V Combine the information above with data on the abundance of FLB and the abundance of the natural bacterial assemblage to calculate the total number of bacteria c o n s u m e d per protist.
B FLB
DISAPPEARANCE
Remove and preserve samples i m m e d i a t e l y and at specific time intervals.
Calculate the rate of c o n s u m p t i o n of natural bacteria in the incubated sample from the rate of disappearance of FLB and the ratio of natural bacteria to FLB.
Microscopically determine the abundance of FLB remaining in the sample.
"o >,
Figure 15.3. General scheme for measuring feeding rates of bacterivorous protists via (A) the uptake of fluorescently labeled bacteria (FLB), and (B) the disappearance of FLB.
Equipment
and reagents ° m
• Container for growing bacteria (e.g. 2 I Fernbach flask); bacteriological growth medium; assorted glassware for transfer and holding seawater and bacterial concentrates. • Cultured bacteria, or concentrate of natural bacterial assemblage (for the latter, see the method of Sherr et al., 1987). 303
0.
1.2
(a)
~0.8~ 0.6o ~ 0.4~0.201 0
50
100 150 Time (minutes}
200
250
2.2 (b)
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5
10
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15 Time (days}
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25
30
Figure 15.4. Examples of results using FLB to measure the feeding rates of bacterivorous protists. (a) Uptake of FLB by two species of antarctic choanoflagellares that differ in size and feeding rate. (b) Disappearance of FLB in a water sample collected from Great South Bay, Long Island.
DTAF (5-(4,6-dichlorotriazin-2-yl) aminofluorescein). Phosphate-saline solution (0.05 M Na~HPO4-NaCI). 0.02 H pyrophosphate-NaCI solution. High-speed centrifuge with rotor for 200 ml polycarbonate bottles (or similar equipment). Sonicator with microtip; vortex mixer. DAPI for bacterial staining and counterstaining of protists (Sherr et al., 1993). Filtration setup; vacuum pump; filtration manifold; 25 mm filtration apparatus (fritted glass base, tower, clamps). Incubation and sampling bottles; e.g. I clear polycarbonate bottles. 304
• • •
•
•
Experimental
Suitable incubation conditions (in situ deployment, or outside incubator at ambient water temperature, as noted in dilution method above). Micropipettes (200 IJl and 1000 lul). Preservative. A 10% glutaraldehyde solution made from a 50% aqueous solution with natural seawater, filtered, and added 1:9 with the sample is often sufficient. However, other fixatives can be used (see item 3 below in 'FLB uptake experiments'). Epifluorescence microscope, equipped with blue light excitation (e.g. Zeiss BP450-490 excitation filter, with FT510 beamsplitter and LP520 barrier filter, or equivalent set) and with UV excitation (e.g. Zeiss BP365 excitation filter with FT420 beam splitter and LP418 barrier filter, or equivalent set). Microscope slides, coverslips, low-fluorescence immersion oil, blackened 0.2 lum and 0.8 lum pore size polycarbonate filters (e.g. Poretics~'). setup
The preparation of FLB from natural bacteria has been previously described (Sherr et al., 1987; Sherr and Sherr, 1993). The published procedure can be followed exactly, or with modifications as noted below. It is best to use as small a bacterial species as possible when using cultured strains because most tend to form large cells in rich growth media relative to the size of bacteria in natural assemblages. Some strains can be starved after growth to reduce their size, but this process often makes the cells excessively 'sticky' and large clumps often form in the cultures. If the strain forms aggregates in the stationary growth phase, then harvest the culture during exponential growth. Grow bacteria in 1 1 of an organic-rich seawater medium, and harvest by centrifugation (~20 000g; 20 min in 200 polycarbonate centrifugation bottles works well, but other volumes and centrifugation schemes can be used). Decant the supernatant, add =50 ml of filtered seawater to each bottle, and vigorously resuspend and disperse the pellets using a vortex mixer. (Note: A phosphate-saline solution can be used for resuspending the cells (Sherr and Sherr, 1993). This can help prevent clumping but is not necessary with some bacterial strains. If used, the saline content of the solution should be matched to the salinity of the water sample.) Once the pellets are dispersed, fill the centrifugation bottles with filtered seawater. Repeat centrifugation and rinse process. Centrifuge and combine bacterial concentrates into a single bottle, and bring the volume to 100 ml with filtered seawater or phosphate-saline solution. Add 20 mg of DTAF and stain for 2 h at 60°C. Then centrifuge and rinse cells three times as described above to remove excess DTAE The final resuspension should be done in 0.02 M pyrophosphate-NaC1 solution (pH = 9) to minimize clumping during storage, unless it is previously established that clumping is not a problem. In some cases, final resuspension and storage in sterile deionized water is adequate. Verify that the cells are adequately stained using an epifluorescence microscope with blue light excitation. Determine and record the abundance of FLB in the final suspension. Dispense the FLB into suitable aliquots (e.g. 1 ml in epitubes). Keep frozen and in the dark until use. 305
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Water samples can be collected and held for experimentation in a variety of containers, but acid-washed, clear polycarbonate bottles are recommended for incubations. Collection and handling should be done aseptically and with as little disturbance as possible (i.e. limit turbulence, bubbling). It is important to maintain natural light regimes and ambient water temperatures during incubations because rates of bacterivory by phagotrophic protists can be highly responsive to light and temperature. Prior to conducting either an FLB uptake or disappearance experiment, a sample should be removed and preserved for enumeration of the natural bacterial assemblage. It is particularly important for FLB uptake experiments to have some knowledge of bacterial abundance in the water. This value is used to determine the number of FLB that will be added to the samples. Also, the natural bacterial abundance is necessary in the calculation of rates of bacterivory in both uptake and disappearance experiments. Determine bacterial abundance by epifluorescence microscopy on blackened 0.2~m pore-size polycarbonate filters. Several methods are available (Kemp et al., 1993). Staining with 4',6-diamidino-2-phenylindole (DAPI) at a final stain concentration of 25-50 t2g ml 1 is recommended because it is the same procedure used to counterstain small protists in the uptake experiments.
FLB uptake experiments 1. Calculate the volume of the FLB concentrate to be added to each sample. For uptake experiments it is necessary to add an appropriate concentration of FLB such that the uptake rate can be determined for the population or community in a reasonable time frame (i.e. before FLB are digested or egested) but less than an amount that will significantly alter the total abundance of bacteria. In practice, FLB are typically added at 5% to 30% of the number of natural bacteria. 2. Dispense water into the incubation bottles (at least triplicates). Thaw a tube of FLB. Vortex the tube vigorously, sonicate (a few seconds at --30 W, using a microtip), vortex again and immediately withdraw the appropriate volume with a 200 1J1 or 1000 1Jl micropipette. The appropriate volume is dependent on the sample volume incubated and the concentration of FLB in the stock. If the concentration of FLB is extremely high in the concentrate, a dilution can be prepared in filtered water and used to inoculate the water sample. In some cases this can aid dispersion of the FLB. Add the FLB to the water samples and mix by gently inverting the bottles. 3. Remove a sample (10-20 ml) immediately and preserve. Record the time of preservation accurately (i.e. elapsed time from the moment of FLB addition). Repeat the sampling at selected times (usually 5-7 samples over a 1-2 h period, depending on the taxa, bacterial concentration and enviromnental conditions). Preservation of samples is critical in FLB uptake experiments because it is imperative that ingested FLB not be egested by the
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phagotrophic protists. Release of food vacuole contents upon preservation is not uncommon among small phagotrophic protists. A variety of preservatives have been used to reduce this problem. In some cases, samples can be preserved by adding filtered, 10% glutaraldehyde (prepared with water from the study site and kept cold) 1:9 with sample to yield a 1% final preservative concentration. Alternatively, a final concentration of 0.5% alkaline Lugol's solution, followed immediately by 2~7~buffered formalin and one drop of 3% sodium thiosulfate can be used (Sherr and Sherr, 1993), or a volume of ice-cold 4(~ glutaraldehyde equal to the sample volume (Sanders, 1988; Bennett et al., 1990). If possible, preliminary work should be conducted to examine the efficacy of these different preservatives for the species/community under study. Place preserved samples in the cold and dark and prepare slides as soon as possible. 4. Examine each time point sample by epifluorescence microscopy at high magnification using oil immersion objectives. Subsamples of each preserved sample are prepared by filtering them onto blackened 0.8 blm pore-size polycarbonate filters. These preparations are counterstained with a final stain concentration of 25-50 pg ml ~DAPI to visualize protists using UV excitation. As each protist is located, switching to blue light excitation allows the number of ingested FLB (if any) to be determined and recorded. Care must be taken to avoid counting FLB that are lying near or on top of protists but not actually ingested.
FLB disappearance experiments Sample and FLB preparation are the same as for uptake experiments (previous section) with one exception. A set of 'control' bottles (bottles containing 0.2 lJm filtered water are inoculated with the same concentration of FLB as the experimental bottles and incubated alongside the experimental bottles. Changes in the FLB concentration in the control bottles are indicative of non-grazing mediated losses (e.g. fading, adherence to the walls) that must be accounted for in long incubations used in this approach.
1. Calculate the volume of the FLB concentrate to be added to each sample. Unlike FLB uptake experiments, the abundance of FLB in disappearance experiments can be a small percentage of the abundance of natural bacteria (typically _<10% of the abundance of natural bacteria) because FLB abundance in the water sample can be assessed more readily than FLB uptake into individual protists. 2. Remove and preserve a sample after FLB are added and mixed into each water sample. Record the time. Preservation is much less critical for FLB disappearance experiments because one need only stop grazing activity and prevent any loss of fluorescence of tile remaining FLB. Continue to remove and preserve samples periodicially over an appropriate time period (usually 1-3 samples over a 12-48 h period). Place samples in the cold and dark and analyze (or freeze) as soon as possible.
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3. Prepare slides for epifluorescence microscopy as for uptake experiments, but do not counterstain with DAPI. Examine slides prepared from each time point (experimental and controls) at high magnification with blue light excitation to determine the total number of FLB remaining at each sampling time. Samples collected at the end of the experiment should also be stained and counted for the number of natural bacteria present (and calculation of the bacteria:FLB ratio at the end of the experiment).
Analyses The abundance of natural bacteria and FEB (and the bacteria:FLB ratio) must be determined in both uptake and disappearance experiments in order to calculate rates of bacterivory in the water sample. For FEB uptake experiments, the average number of FLB per protist is determined by microscopy on samples collected at each time point. A plot of the average number of FEB per protist versus time generally yields a linear relationship for the initial time points (Figure 15.4(a)), although this relationship may level off as the experiment progresses due to digestion of ingested FEB. It is also important to note that this relationship may not have a Y-intercept equal to zero if some non-ingested FEB were counted as ingested (e.g. if they were laying on top of the protists). The slope of the linear portion of this regression is the rate of FEB ingestion per protist time '. This value is converted to total bacteria ingested per protist time ' by multiplying by the bacteria:FLB ratio for the taxa or group of interest (e.g. heterotrophic flagellates, ciliates). Clearance rates for these taxa are determined by dividing the FEB ingestion rates (FEB per protist time 1)by the concentration of FEB. These values are usually expressed in I,ll per protist hour'). Rough estimates of community-level bacterivory are obtained by multiplying group-specific clearance rates by the abundance of each group (the latter determined using standard microscopical methods (Kemp et al., 1993)). Estimates of community-level bacterivory obtained in this way are accurate only if the microscopical examination adequately assesses all bacterivorous species. For FLB disappearance experiments, the abundance of FEB at each time point is plotted against time of incubation (after accounting for any non-grazing losses of FLB in the controls). The community-level rate of bacterivory is calculated directly from this rate of disappearance of FEB (Figure 15.4(b)). Two important considerations for calculating the rate of FEB disappearance from these data are (1) the choice of a linear or exponential model for the rate of disappearance and, (2) changes in the ratio of bacteria:FEB during the incubation. The choice of a mathematical model can be difficult if only 'endpoint' determinations of FEB abundance are made, and if natural bacterial abundance is determined only at the beginning of the experiment. Examination of the time course plots of FEB vs. time can aid the former decision (see Figure 15.4(b)), while changes in the bacteria:FEB ratio can be addressed by direct counts of FEB and natural bacteria at the beginning and end of the incubation 308
period. Suitable mathematical models can be found in Salat and Marras6 (1994).
Caveats, precautions and alternatives A variety of surrogate prey have been employed to examine rates of bacterivory in natural water samples. These include fluorescent latex microspheres (Borsheim, 1984; Cynar and Sieburth, 1986), fluorescent paint particles (McManus and Fuhrman, 1986) and genetically marked minicells of Escherichia coli (Wikner et al., 1986; Wikner, 1993). These various methods have specific advantages, but they also have distinct disadvantages that range from complicated/radioactive protocols to feeding selectivity against surrogate prey. The uptake a n d / o r disappearance of FLB is a versatile and relatively straightforward method for examining rates of bacterivory in natural microbial assemblages. An advantage of the method is that natural bacteria are used as prey to examine bacterivory, rather than latex or other materials which may be selected against by some microbial bacterivores (Sherr et al., 1987). The method has the additional advantage that it can be used to examine rates of bacterial ingestion by specific microbial taxa or groups (by FLB uptake), or community-level bacterivory (by FLB disappearance). Semi-automated determinations may further improve the approach. Analysis of FLB by flow cytometry can be relatively straightforward for FLB disappearance experiments, and the instrument can also be used to determine the abundance of natural bacteria (del Giorgio et al., 1996). Methodology for the application of flow cytometry to FLB uptake experiments has recently been published (Bratvold et al., 2000). Finally, DTAF has been used effectively to stain eukaryotic algae (Rublee and Gallegos, 1989), allowing analysis of microbial herbivory. Nevertheless, there are some important caveats to the method. It is clear that prey size is an important criterion for particle ingestion by phagotrophic protists (Gonz~lez et al., 1990; Simek and Chrzanowski, 1992). FLB populations rarely mimic the size spectrum of natural bacterial assemblages. In addition, chemical aspects of microbial species also affect their acceptability as prey. Experimental studies have indicated that live bacteria may be preferred to heat-killed cells (Landry et al., 1991). These selective grazing behaviors may affect the ingestion of FLB relative to unlabeled cells. The use of vital stains for producing FLB may alleviate some of these issues (Epstein and Rossel, 1995) but the appropriateness of FLB will continue to be questioned. Two important issues that relate to FLB uptake experiments are (1) the number of FLB added to samples and, (2) the potential for the release of FLB. The number of FLB added to a water sample must be a compromise. Enough FLB must be added to allow them to be taken up frequently by phagotrophic protists (to facilitate microscopical documentation), but at concentrations that will not substantially alter the bacterial abundance in the sample. Significant increases in total bacterial abundance can increase ingestion rates because of the increased availability of prey. Ideally, FLB
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should not be added at concentrations greater than 30% of the abundance of the natural bacterial assemblage, but values up to 50% are not uncommon. Egestion of FLB in uptake experiments is a major concern. Some protists evacuate food vacuoles when preservatives are added to samples, resulting in underestimations of their FLB uptake rates. The use of icecold preservatives (Bennett et al., 1990) or more toxic concoctions may help prevent egestion (Sieracki et al., 1987) but it is never assured that all egestion has been prevented. One important topic relating to FLB disappearance experiments is the fading of FLB in long-term (e.g. 24 h) experiments in high light or warm water environments. Microscopical counts of faded FLB can be difficult, and it is imperative to perform counts on control treatments (FLB in filtered water) incubated alongside experimental bottles. Decreases in the concentration of FLB that are unrelated to grazing-mediated disappearance can reduce the sensitivity of this approach.
CONCLUSIONS AND RECOMMENDATIONS The experimental protocols described here to measure microbial bacterivory and herbivory are two of many approaches and variations that have been employed to investigate these processes. As stated previously, no single method is foolproof, and none is appropriate in all ecosystems and circumstances. Given this uncertainty, it is advisable to use more than a single method whenever possible. It is particularly prudent to try to link production and removal of the same assemblage. For example, it is logical that measurements of bacterial production and bacterivory should be reconcilable with net changes in bacterial abundance in an ecosystem. If production exceeds removal, net changes should be positive, while if removal exceeds production, net changes should be negative. A three-way comparison of this type in the Sargasso Sea yielded useful insights into potential artefacts within the different methological approaches (Caron et al., 1999). Similarly, comparisons of phytoplankton growth and mortality have yielded information that is pertinent to the prediction and understanding of phytoplankton blooms and the vertical flux of primary production (Landry et al., 1998; Caron and Dennett, 1999). Eventually, it would be highly desirable to move towards methods that do not rely on the incubation of microbial assemblages to make measurements of growth and grazing rates. Some early attempts in this direction are promising (GonzSlez et al., 1993), but thus far the interpretation of those protocols must still be tied to techniques that depend on incubation. Nevertheless, these innovative methods may offer the ability to augment more traditional approaches because numerous analyses can typically be performed using these 'snapshot' techniques. The bottom line is that an appropriate choice of methodology is predicated on a reasonably good knowledge of the system(s) to be studied, and a thorough understanding of the advantages and limitations of each 310
technique. As a c o n s e q u e n c e , there is typically a significant l e a r n i n g c u r v e for this t y p e of e x p e r i m e n t a l w o r k .
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16 Flow Cytometric Analysis of Autotrophic Picoplankton L Campbell Department of Oceanography, TexasA&M University,College Station, TX 77843-3146, USA
CONTENTS Introduction Methods Analysis and interpretation of results Discussion and future directions Resources
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INTRODUCTION Flow cytometry is a valuable, if not essential, tool for studies of aquatic microbial populations. The technology of flow cytometry permits the simultaneous measurement of multiple properties of individual cells in suspension. Although flow cytometry was developed for mammalian cell systems, the capabilities of this instrumentation for rapid enumeration and quantification of both structural and functional properties of individual cells makes it ideal for applications in marine microbial ecology. The enormous potential of flow cytometry in the study of marine microbial ecology is well recognized. Since the early 1980s (Yentsch et al., 1983; Olson et al., 1983), the number of flow cytometry applications in marine microbial ecology has increased tremendously (for reviews see Olson et al., 1991; Fouchet et al., 1993; Porter, 1999; Collier and Campbell, 1999). Flow cytometry has become an indispensable tool in marine microbial ecology for the identification and enumeration of picoplankton, that is, cells that are < 2 pm in diameter (Chisholm et al., 1986). The recognition of the importance of picoplankton has resulted in a fundamental change in our understanding of the food web of the ocean (Pomeroy, 1974; Azam et al., 1983). An important goal in microbial ecology is to increase our understanding of the roles individual organisms play in oceanic food webs. To do this we will employ the rapidly increasing number of tools we have available to collect information on activities at the single-cell level. The aim of this chapter is to provide an introduction for both the experienced flow cytometrist faced with the challenge of investigating cells
METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0-12 521530 4
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less than 2 l~m in diameter and the microbial ecologist who wishes to begin flow analyses. A brief introduction includes an overview of the principles of flow cytometry, with references to some of the important reviews of methodology and applications. Comments are included throughout the Protocols for consideration when applying this basic protocol in new applications. The focus of this chapter is on analysis of autotrophic picoplankton using a popular benchtop instrument, the FACSCalibur (Becton Dickinson). Note also that the focus is on analysis rather than sorting. Although sorting is one of the most powerful uses of flow cytometry (Yentsch et al., 1983; Olson et al., 1993; Robinson et al., 1997), the specifics are largely instrument-dependent and are not amenable to description in a general protocol. Finally, there are many excellent resources available that are useful for both the novice and experienced investigator.
Principles of flow c y t o m e t r y Instrumentation Flow cytometry is the analysis of light scatter and fluorescence emitted from individual cells as they flow individually though an intensely focused light source at a rate of thousands of cells per second. To accomplish this, a cytometer consists of three components: fluidics, optics, and electronics for detection and data acquisition. The fluidics system includes a pressurized fluid delivery system, or sheath. The sample is injected into the sheath such that the suspension of single cells is hydrodynamically focused to insure cells are in laminar flow and aligned in single file along the narrow central core through the sensing region. The optical component includes a light source (most often a laser) and a series of dichroic mirrors and bandpass filters to divide the emitted light. The light source must be very focused and intense to provide a large number of photons within the sensing region (beam spot) because cells remain within this region only just a few microseconds. As cells pass through the sensing region, a light scatter pulse is generated and collected at both small forward angles (FALS) and at right angles to the cell (side scatter, SSC). The FALS signal is influenced principally by cell size, whereas SSC is also influenced by refractive index and internal cell structure. The typical fluorescence parameters also collected are the green (GRFL), orange (ORFL), red (RFL) regions, as defined by the optical filters used. An example of the standard optical filter array used to define each parameter in a FACSCalibur (488 nm laser) is depicted in Figure 16.1. In the detection system, the light pulse is converted to an electrical signal (voltage) by either a photomultiplier (PMT) or photodiode. Because side scatter and fluorescence are usually very weak signals, PMTs are used to amplify the signals. The FALS signal is stronger, so is most frequently collected by a photodiode. Using a PMT for FALS detection, however, permits greater sensitivity by improving the signal to noise ratio, and some instruments have been modified to detect FALS using a
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Figure 16.1. Standard optics layout for the FACSCalibur flow cytometer. A cell intercepts the focused laser beam within the sensing region of the flow cell. Light is scattered by the cell in the forward angle light and detected by a photodiode (FALS). Light scattered at right angles to the cell passes through a dichroic filter that splits the light into wavelengths > 560 nm and < 560 nm. Fluorescence > 560 nm is subsequently split again with a 640-nm-long pass filter. The wavelengths > 650 nm are detected by the red fluorescence (RFL) PMT and the range of wavelengths within approximately 560-640 nm is detected by the orange fluorescence (ORFL) PMT. Fluorescence within the range 515-545 nm is collected by the green fluorescence (GRFL) PMT.
PMT ( D u s e n b e r r y and Frankel, 1994). The high voltage (HV) on each PMT can be adjusted to optimize signal amplification. Logarithmic amplification is u s e d m o s t frequently to p r o v i d e a w i d e d y n a m i c range, often four decades. The analog signal is then converted to a digital value. In m o s t m o d e r n flow cytometers, the signal range is converted into 1024 channels (256 channels per decade). Thus, each r a w data value is stored as a channel value. Most flow cytometers are capable of collecting and recording at least five p a r a m e t e r s for each cell. Data are stored in listmode files, in which each event with the c o r r e s p o n d i n g data for each p a r a m e t e r is recorded sequentially.
319
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Analysis The strategy in analysis of flow cytometric data is to distinguish individual unique populations among the acquired events. Many analyses are based on principal component or clustering analysis (Watson, 1992; Wilkins et al., 1994) and the newest approaches utilize neural networks (Frankel et al., 1989, 1996; Smits et al., 1992; Davey et al., 1999). Data are generally plotted in a series of two-parameter histograms to identify specific populations. Multiparametric analysis makes it possible to characterize populations of cells based on correlating parameters of cell size, shape, or granularity of populations, as well as fluorescence color and intensity. There are a number of fluorescent stains that are useful in flow cytometry because they react with specific cellular components, such as DNA content and total protein, or allow properties such as membrane permeability to be determined (Porter, 1999). The photosynthetic pigments of phytoplankton provide a natural fluorescence marker for both identification and to infer pigment content (Yentsch et al., 1983; Olson et al., 1991). Fluorescently labeled antibodies are another means of tagging specific populations by surface or cytoplasmic receptors a n d / o r antigens, thereby providing an alternative fluorescence parameter for quantitative analysis (Vrieling and Anderson, 1996; Shapiro and Campbell, 1998). Enzyme activities or intracellular ion levels can also be determined using specific functional probes and fluorophores (Molecular Probes; see Suppliers). Recently, fluorescently labeled oligonucleotide probes have been added to the suite of available tools for identification of specific microbial cells (Amman et al., 1990; Wallner et al., 1997; Simon et al., 2000). A variety of fluorophores are available for use as reporters (see Table 16.1).
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METHODS
Equipment and supplies Equipment Flow cytometer. The choice of instruments for flow cytometric analyses is dependent primarily on the intended applications, as well as the budget available. Although it is possible to build one's own (C)lson et al., 1983; Shapiro, 1995), the majority of instruments used by marine microbiologists are manufactured by Becton Dickinson (FACScan, FACSordFACSCalibur, FACSVantage) and Coulter Electronics (Epics V, 541,753, Elite, and XL) (see Suppliers). In a recent survey of selected oceanographic and environmental microbiological research institutions, the instruments currently in use are largely the newer, benchtop models with air-cooled lasers and optics that require minimal alignment by the operator (see Dubelaar and Jonker, 2000). These relatively compact instruments, although developed primarily for clinical applications, have been adopted by many ecological laboratories because of their relatively simple requirements for operation, the ease with which they
320
can be taken on board ships for oceanographic research, and their improved optics and sensitivity. For analysis of microbial populations, high sensitivity optics are required, in particular for accurate analysis of Prochlorococcus, the extremely small and wealdy fluorescent photosynthetic picoplankter (Chisholm et al., 1988; Olson et al., 1990; see Advantages and limitations). The larger research instruments employed in much of the original oceanographic applications of flow cytometry (e.g. Coulter EPICS instruments; Yentsch et al., 1983; Olson et al., 1989) employ a tunable laser (in some cases two lasers), whereas most of the newer benchtop models are equipped with a single, fixed wavelength 488 nm (blue light), air-cooled laser (15 mW). In some instruments an arc lamp is utilized as an illumination source (Bryte HS, BioRad) or as an additional UV illumination source (Particle Analysis System, Partec GmbH). The newest Becton Dickinson instrument, BD LSR, has an air-cooled UV laser in addition to the 488 nm laser. Along with the improvements in optics, rapid advances in personal computing power have added significantly to the capabilities of flow cytometric analyses. A number of laboratories have upgraded the data acquisition capabilities of older systems by adding a CICERO data acquisition system (Cytomation, Inc.). This product is designed specifically to replace outdated electronics and computer systems while making use of existing optical and fluidics components. None of the commercially available instruments can meet all the requirements desirable for analyses of environmental samples. Given the small market demand it is unlikely that new commercial instrumentation will be developed that incorporates such specifics (Dubelaar and Jonker, 2000). For specialized applications, however, a number of modifications to existing systems, as well as custom-built systems, have been employed to address specific questions. One of the primary challenges for phytoplankton ecologists is the range in cell size and abundances of phytoplankton cells; larger cells are several orders of magnitude less abundant than the smaller picoplankton. In one approach to address this problem, an EPICS 753 (Coulter) was modified with the addition of a dual-sheath to permit varying sample flow rates appropriate for cells of different size and abundances (Cavendar-Bares et al., 1998). Another approach to assess the larger-sized phytoplankton is the EurOPA (European Optical Plankton Analyzer; Dubelaar et al., 1989; Dubelaar and van der Reijden, 1995) which has a total signal range of seven orders of magnitude. The EurOPA was designed to process large volumes of sample, which is important for the analysis of marine samples with low densities of algae. Environmental abundances of most prokaryotic picoplankton, however, generally matches the optimum sample density for flow cytometric analysis (> 10~to 10>cells ml '). Another useful parameter in many flow cytometry applications is the determination of cell volume. The 'Coulter volume,' or impedance volume, was a feature on earlier models (Phinney and Cucci, 1989), but is lacking on most current models of flow cytometers. Given the importance of this parameter in many applications, a customized instrument utilizing
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both optical and electrical (Coulter volume) cell analysis has recently been built by Wietzorrek and co-workers (1999). Given the range of instruments and possible configurations, specific operating instructions will vary among commercial flow cytometers. For the purposes of this chapter it is assumed an experienced operator is available to provide general advice on instrument operation. • • •
Ultrafreezer (-80°C) for sample storage Liquid nitrogen dewar for quickly freezing samples Networking capability for data transfer
Supplies • • • •
Sample tubes for analysis of appropriate size for instrument Pipettes (for 10 tJl to 1000 pl volumes) 0.22 lum sterile filter units (Acrodisc) Large volume 0.22 pm filter apparatus (cartridges or glass apparatus; autoclavable) • Cryovials for samples preserved and frozen; or dark plastic bottles for storage of live samples • Vortexer • Data archival tape or CD-ROM
Solutions • Preservative. If samples cannot be processed live, they must be preserved and stored such that cellular structure and fluorescence is maintained. Both glutaraldehyde (Vaulot et al., 1989) and paraformaldehyde (0.2% final concentration, Campbell et al., 1994; or I% final concentration, Marie et al., 1999b) have been used successfully. Preparation of 10% paraformaldehyde stock I. 2. 3.
4. 5. 6. • •
Weigh 2 g paraformaldehyde. (Use caution! Weigh in hood with minimal air turbulence.) Add 18 ml boiling ddH20 and stir to dissolve. Do not continue to heat. Add I N NaOH drop-wise (only a few drops will be required for this small volume) and continue stirring until precipitate dissolves completely; this may take an hour or longer if a larger volume of the stock solution is prepared. Cool to room temperature. Add 2 ml of 0.22-1am-filtered seawater, or 0.02 M phosphate-buffered saline (PBS) buffer, and adjust pH to 7.4. Filter-sterilize using a 0.22 lure filter disk (Acrodisc, Gelman) directly into a sterile tube and store at 4°C. Use within I week.
Liquid nitrogen Sheath. Ideally, the sheath fluid should be of the same refractive index as the samples to provide the most accurate analysis.Thus, filtered seawater of the same salinity as the samples is optimum. In practice, distilled deionized water
322
(ddH20) can be used as sheath for most analyses of marine picoplankton. If, however, cells are to be sorted for establishing cultures or for subsequent analyses, seawater (or artificial seawater) must be used to maintain cell integrity. Sheath must be particle-free, so must be filtered through 0.22 lure filter immediately prior to use.The in-line filter included in the fluidic system of most (all) flow cytometers is therefore critical and must be replaced frequently if usage is high or if blanks (filtered seawater controls) are not acceptable. • Calibration beads. Commercially available polystyrene beads of uniform size containing fluorophores with fluorescence properties similar to phytoplankton pigments are useful for both verification of instrument set-up and are necessary for normalizing cellular fluorescence properties of field populations (see Analysis). Examples of useful beads include Yellow-Green Fluoresbrite (Polysciences, PA) or Purple-Yellow (Spherotech, Libertyville, IL) fluorescent beads in the I-2 lum diameter range. Working stock dilutions of calibration beads are prepared by diluting original stocks with particle-free ddH20 and preserving with azide (0.02% final concentration). Vortex stocks to reduce clumping and doublet formation; however, note that some bead types should not be vortexed. Concentrations of working stocks should be prepared so that by adding 5-10 lul to each I ml of sample the calibration bead concentration will be approximately 104 to I 0s beads ml ~.Thus, there will be several thousand bead events recorded in each analysis. • Stains and fluorophores. For analyses of photosynthetic picoplankton, the autofluorescence of photosynthetic pigments provide natural markers (see Table 16.1). For non-autofluorescent populations, e.g. the heterotrophic bacteria, a nucleic acid specific dye is necessary to distinguish living cells from non-biological particles. Examples include the UV-excitable dye Hoechst 33342 (I lug ml' final concentration) in systems with dual-laser capabilities (Monger and Landry, 1993; Campbell et al., 1994).The advantage of a dual beam technology is the greater spectral separation of UV-excitable nucleic acid stains and chlorophyll fluorescence (see Table 16.1). Newer dyes, excitable with blue light provided by the standard 488 nm laser in the benchtop instruments, are also useful in bacterial analyses (e.g. SYBR, SYTOX, Picogreen; Marie et al., 1997;Troussellier et al., 1999; Veldhuis et al., 1997; see Chapter 8, this volume). However, because of the spillover effect from the green fluorescence of these nucleic acid stains into other fluorescence channels, accurate analyses of the fluorescence parameters of the autotrophic picoplankton can be obtained only from unstained samples. Thus, both unstained and stained replicates should be run if both autotrophic and heterotrophic components of the picoplankton are to be analyzed.Another application of DNA-specific stains is in phytoplankton cell cycle analyses for determination of cell-specific growth rates (seeTable 16. I). Additionally, a variety of reporter fluorophores are now available that can be conjugated to antibodies or oligonucleotide probes for use in flow cytometry (e.g. Shapiro, 1995; Molecular Probes, http://www.probes.com).A summary of some of the commonly used fluorophores is listed in Table 16. I~ New stains should be evaluated for specificity and efficacy in the system in which they are employed.
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Table 16.1 Fluorophores used in flow cytometry and their applications in marine microbial
ecology Fluorophore
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DNA; cell cycle DNA; cell cycle Reporter Reporter Reporter
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Reporter FITC: protein stain, Reporter Reporter Reporter DNA; cell cycle DNA DNA, RNA DNA; live/dead; cell cycle Reporter
(1) (2)
For UV-laser excitation
Hoechst 33342 DAPI Cascade Blue AMCA Alexa350
Blue Blue Blue Blue Blue
For 488 nm excitation
Cy2 Fluorescein
Green Green
489 494
502 518
FluorX Alexa488 SYBR PicoGreen SYTO13 SYTOX Green
Green Green Green Green Green Green
494 495 494 502 488 504
520 519 521 523 509 523
PE (phycoerythrin)* Cy3 Alexa546 SYTOX Orange
Orange
545
575
Orange Orange Orange
550 556 547
570 575 570
Red Red
490 518
675 605
Red
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Red Red
589 590 649 642 650
PerCP Ethidium bromide Propidium iodide Texas Red Alexa594
Reporter Reporter DNA; live/dead; cell cycle Reporter DNA, RNA
(2) (2) (2) (2) (2) (2)
(1) (2) (2) (2) (2) (2)
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DNA, RNA, dead-cell stain Reporter Reporter
670 661 660
Reporter Nucleic acids Reporter
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For 633 nnl excitation
Cy5 Red TOTO-3 Red APC Red (allophycocyanin)
(1) Amersham l~harmacia Biotech; (2) Molecular Probes. * Cyanobacterial photosynthetic pigment; the diagnostic fluorescence parameters for the photosynthetic picoplankton include RFL due to chk~rophyll a (absorption maxima at 43t) nnl and 662 nm; emission maximuna between 080 688 nm), which is present in all algae and cyanobacteria, and ORFL due to phycoerythrin, which is present in the marine cyanobacteria Syiluch~coccus spp. Both pigments are exci~ed by 488 nm light.
324
Protocols Enumeration of autotrophic picoplankton Sample collection and preservation Samples collected in the field with Niskin bottles, or from laboratory culture experiments, should be protected from direct sunlight prior to analysis. To preserve sarnples for analysis on return to the laboratory, 201Jl paraformaldehyde stock (see Solutions) is added per ml of sample in a cryovial. After fixation at room temperature for 10min samples are quickly frozen in liquid nitrogen. Samples should be stored at -80°C to preserve fluorescence (Jeffrey et al., 1997). Replicate 1 ml samples should be collected. Prepare sheath and particle-free water for dilutions Approximately 1 1 sheath volume per hour is required during a typical analysis. Particle-free (0.22 tJm filtrate) sheath is necessary for analyses of bacteria. Autoclaved solutions, while sterile, may not be particle-free, so solutions should be filtered immediately prior to use. Prepare particle-free ddH~O (filter twice through 0.22 t_lm disposable filters) for preparation of working dilutions of calibration beads. If analyzing samples from culture experiments, prepare particle-free filtered seawater for sample dilution, as needed (see Analysis). Instrument setup Included here is a basic procedure intended for users familiar with a benchtop model with an air-cooled laser. For additional information see Olson at al. (1989, 1993), and see Robinson (1993) for sample start-up procedures for a variety of instruments. 1. Fill sheath tank; empty waste tank. 2. Turn on power; allow laser to w a r m up. 3. Verification of alignment and setup. The optical filter array (e.g. FACSCalibur standard optical setup, Figure 16.1) is appropriate for analysis of autotrophic picoplankton. Optical filters m a y not be identical among instruments. So if different instruments are used, verify that the dichroic mirror or bandpass cutoffs are compatible with your application. Establish initial conditions Background noise (number of events recorded} should be minimal; this is verified by recording the event rate while running a sample tube with sheath or ddH20 only.The optical path and flow cell must be clean and free from dust; this is especially important in analyses of picoplankton, in which noise can be troublesome. • Verify that the calibration beads match previous cytograms under standard conditions To verify that the instrument is operating normally, load standard
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settings (as above) and a template that includes a bead 'window' (see Figure 16.2). Calibration beads should appear as a tight population and be located in the same position relative to the bitmap or 'window' in the displayed histogram.The coefficient of variation (CV) for commercially prepared fluorescent calibration beads is provided by the manufacturer for each parameter (e.g. I-3%, Spherotech, Inc.). An example of calibration beads run on a FACSCalibur instrument is shown in Figure 16.2.
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Figure 16.2. Cytogram template for Purple:Yellow Calibration beads (2.0 pm, Spherotech, Inc.). Bivariate histogram of red fluorescence (RFL) vs. forward angle light scatter (FALS) and the corresponding one-parameter histograms for each parameter are plottbd~'The coefficients of variation (CV = standard deviation/ mean x 100) for linearized data for each parameter are the following: FALS (CV= 3.0%); SSC (CV = 8.7%); GRFL (CV = 4.6%); ORFL (CV = 5.2%) and RFL (CV = 9.1%). Data were plotted using WinMDI (J. Trotter, San Diego, CA) and Powerpoint (Microsoft Corp.).
Analysisof autotrophicpicoplankton Once the i n s t r u m e n t operation has been verified, proceed with s a m p l e analysis. Note that once they are thawed, samples cannot be re-frozen, and generally cannot be stored m o r e than a few hours at 4°C, so it is p r u d e n t to establish n o r m a l operation before t h a w i n g samples. 1. T h a w frozen s a m p l e s in running tap w a t e r and store in d a r k and on ice if not analyzed immediately. 2. A d d internal calibration b e a d s (e.g. 5 pl) to s a m p l e tube followed by 0.5 ml s a m p l e and t h o r o u g h l y mix sample.
326
If samples are from coastal waters or if known to contain large aggregates, samples should be pre-filtered through Nitex mesh to avoid clogging the flow cell (see Discussion). 3. Load appropriate settings and acquisition template for field samples. Set amplification to log scale. For cell cycle analysis, when DNA fluorescence must be collected on a linear scale (Boucher et al., 1991; Marie et al., 1997) linear amplification instead of log is used. Create a new data file folder on your computer, and set a unique file prefix for files to be saved during this run (note, the date is often the most useful file prefix). Prior to running environmental samples, it is useful to run cultures of known cell types so that the range in gain settings for the FALS detector or HV settings for SSC and fluorescence PMTs can be approximated. Once appropriate HV settings are known, they can be saved as a template file. For example, typical settings for analysis of an oceanic picoplankton sample using a FACSCalibur instrument would be: FALS = E01; SSC = 450; GFL = 600; ORFL -- 600; RFL -600. Set threshold parameter on SSC starting at channel 52, or higher if necessary to eliminate noise. Alternatively, threshold can be set on RFL. Most modern flow cytometers have a four-decade log scale so the range of picoplankton populations will be on scale simultaneously. To enumerate the larger cells, which may be off-scale at standard picoplankton settings, decrease HV setting and run the sample a second time. However, in many cases, the larger cells will be present at much lower densities, so must be run at increased flow rate (see Olson et al., 1991; Cavendar-Bares et al., 1998). Instruments such as the EurOPA (see above) have alternative strategies for obtaining counts over a dynamic range of cell sizes. 4. Enter sample identification number and information in your data log sheet with corresponding instrument run number. 5. Place sample on sample introduction port and allow system to return to operating pressure before beginning data acquisition. Field samples should be run at the highest flow rate, e.g. 60 ~1 min ', whereas cultures may be run at lower flow rates (see Calibration). The voltage indicating flow rate is shown on the status menu and should be monitored carefully before running each new sample. 6. Verify that HV and gain settings are optimized for the sample. Note, the HV for the RFL PMT is set higher for samples from surface mixed layer waters (where cells have less pigment per cell) than for deeper samples (Figure 16.3). 7. Acquire data using a fixed time for acquisition. For accuracy, thousands (10 000) of events for each population, rather than total events, should be acquired. Thus, run time will depend on cell abundance for each population of interest. Typically 2 to 4 min acquisition at 60 pl min 1 is required for field samples from the oligotrophic ocean. For cultures, 30-60 s is often sufficient; however, culture samples may require dilution to obtain a reasonable event rate (<1000) as well (see Analysis), 327
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8. A control, or filtered sea water blank, should always be run. A blank is prepared by pipetting sheath or d d H , O into a sample tube and running at the standard HV and gain settings used for analysis. 9. Save data to a listmode file. For picoplankton analysis, the parameters saved to listmode file should include: FALS, SSC, GRFL, ORFL, RFL and time (duration of run). 10. At completion of sample analysis, run ddH~O to rinse seawater from the system. Next, run 10% bleach (5 min) followed by d d H , O (5 min), which is the standard r e c o m m e n d e d daily cleaning protocol. At m o n t h l y intervals, remove the in-line sheath filter, and clean the entire system by running 10e7, bleach through both the sheath and sample lines using a sheath tank dedicated to bleaching the system. If seawater is used as sheath, the sheath system needs to be thoroughly rinsed with ddH~O daily. Replace seawater sheath tank with ddH~O sheath tank and flush system for 30 to 60 min. 11. S h u t d o w n procedure. Following the manufacturer's instructions, allow laser to cool and then shut off power.
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Data analysis software Listmode data can be processed directly using manufacturers' software (e.g. CellQuesff M Becton Dickinson; CYCLOPS TM, Cytomation; Elite TM, Beckman Coulter). Commercially developed software (Verity, D e N o v o Software, Phoenix Flow) and freeware (Win MDI, J. Trotter; CYTOWin, D. Vaulot) are also available. As an example, CYTOWin is a Windows-based version of CYTOPC (Vaulot, 1989) to plot and analyze flow cytometric listmode files (available to the scientific c o m m u n i t y at http://www.sb-roscoff.fr/Phyto/ cyto.html). The user can define up to nine w i n d o w s or bitmaps, and the n u m b e r of events, mean, m o d e and coefficient of variation are determined for each window. The logarithmic to linear transformations for each parameter are p e r f o r m e d automatically based on coefficients supplied by the user (see Calculation) and each parameter can be normalized to a specified calibration bead fluorescence. The strategy in the analysis of flow cytometric data for picoplankton is to distinguish individual populations and to quantify the abundance and optical properties for each. In Figure 16.4, a set of histograms is used to demonstrate h o w each of the autotrophic picoplankton groups
Figure 16.3. Example of the variation in cellular chlorophyll fluorescence (RFL per cell) for autotrophic picoplankton in field samples collected from 25, 70, 90 and 130 m at a station in the South Pacific (18~ 1' S 177° E) and analyzed live. RFL of the picoplankton populations increases as depth increases; this can be seen by comparing RFL of the cells to the RFL of the calibration beads.
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Figure 16.4. Analysis of a listmode file using CYTOWin (D. Vaulot, Roscoff, France) to enumerate autotrophic picoplankton, Prochlorococcus, Synechococcus, and the small eukaryotic algae. The three picoplankton populations can be distinguished based on scatter (size) and fluorescence (color and intensity) determined using three bivariate histograms: side-scatter vs. chlorophyll fluorescence (SSC vs. RFL); side-scatter vs. phycoerythrin (orange) fluorescence (SSC vs. ORFL) and chlorophyll vs. phycoerythrin (RFL vs. ORFL).
(Pmchlorococcus, Synechococcus, and small eukaryotic algae) is distinguished in this sample from the oligotrophic South Pacific Ocean. 1. Plot data in a series of bivariate histograms to visualize data: FALS vs. RFL, SSC vs. RFL, RFL vs. ORFL, SSC vs. ORFL. In some cases it is useful to plot RFL vs. GFL or ORFL vs. GFL to distinguish Synechococcus multiple populations. 2. Draw w i n d o w s or bitmaps around each population in each histogram to define the population. Boundaries for each population are determined by inspection. Once all populations are defined, the file can be replayed to verify or adjust windows. 3. Data are saved in an o u t p u t file that can be imported into a spreadsheet file.
Calculation The abundance, N (cell ml '), for each population in a field sample is calculated from the equation:
N= C / ( T x R ) x C F x 10001alml ' where C is the n u m b e r of events acquired (ceils) for a specified population (see Analysis), T is the duration of analysis (min), R is the sample delivery rate ( pl min ' (see Calibration), and CF is a correction factor to account for sample dilution owing to preservation, bead addition, or staining. Typical abundance profiles for picoplankton populations are shown in Figure 16.5. Cellular RFL measurements can be used as an index of chlorophyll per cell, so this parameter is useful w h e n interpreting time-course experiments, diel patterns, or seasonal patterns in time series data. The average
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Chlorophyll (red) fluorescence per cell Figure 16.5. (a) An example of depth profiles of abundance for Prochlorococcus, S~jnechococcl~s, and the small eukaryotic algae. The sample was collected at a station in the Gulf of Mexico at 25° 2Y N 93'-'35' W. (b) Cellular chlorophyll fluorescence (RFL per cell) profiles for each of the picoplankton populations in (a). All values are normalized to P-Y beads (Spherotech, inc.) run simultaneously with the sample.
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~8 cellular fluorescence for each p o p u l a t i o n m u s t be n o r m a l i z e d to a fluorescence calibration b e a d ' s t a n d a r d ' fluorescence to a c c o u n t for differences in i n s t r u m e n t settings a m o n g a n a l y s e s (see Figures 16.3 a n d 16.5). FALS m e a s u r e m e n t s can be u s e d to estimate cell size. Size is m o s t directly related to f o r w a r d angle light scatter, h o w e v e r , this relationship m u s t be calibrated ( S h a l a p y o n o k et al., 2000).
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Note that all data collected in log scale must be converted to linear values before computing statistical analyses. If X = linear channel n u m b e r and Y -- log channel number, X = a Y ~'. Many software packages assume a = 1 and b -- 4, but the actual coefficients are amplifier specific and must be calibrated (Vaulot, 1989). The log to linear conversion for each PMT is calibrated by comparing fluorescence signals collected on both log and linear scales over a range of HV settings, and then computing the regression of log X and Y. Alternatively, a series of fluorescent beads with k n o w n relative intensities (e.g. Rainbow Beads; Spherotech, Inc.) can be used (see Bagwell et al., 1989; Wood, 1997).
Calibration To obtain accurate cell abundance data, the v o l u m e analyzed must be known. For systems such as the FACSCalibur, the v o l u m e analyzed is determined from Volume = flow rate x time It is therefore important to calibrate the sample flow rate (delivery rate) accurately. This is most simply accomplished by the procedure recomm e n d e d in the BD manual and Marie et al. (1999b): 1. Weigh a sample tube with 1 ml of water to determine initial v o l u m e (Vi) = weight x density. 2. Remove the outer sleeve of the sample injection tube to prevent sample aspiration by a droplet retrieval system. 3. Place tube on introduction p o r t - - d o not allow a drop to fall into tube. 4. Run water at HI setting for precisely 10 rain. 5. Weigh remaining water v o l u m e to calculate final (V~) and determine the v o l u m e (ml) run. 6. Rate = (VI- V~) / 10 rain x density of water. Freshwater density at room temperature is 1.0 g cm ~. Use the rate determined in (6) to calculate abundance (see Calculation). At frequent intervals, the flow rate calibration should be verified. Note, the calculated flow rate often does not equal the manufacturer's listed rate (e.g. 60 t21 min ~). As mentioned previously, for analysis using the FACSCalibur, a stable flow rate must be established before beginning data acquisition because the flow rate is d e p e n d e n t on system pressure (determined by the voltage setting). It is very important to allow the system to stabilize so that flow rate will be constant during data acquisition. A second factor that influences the accuracy of calculated cell abundances is the event rate, or count rate. Efficiency of data acquisition will vary with event rate due to coincidence. At high event rates (when cell abundances greater than 10~'-107 cells ml 1), the electronic 'dead time' due to signal processing can become significant, and, consequently, events are 'lost' because processing cannot keep up with the n u m b e r of events. For the FACSCalibur, we have found that counts are accurate if the event rate
332
< 1500. Although a correction factor can be determined for event rates > 1500, it is recommended that all sample analyses be acquired at a rate of below 1000 events s ~ for the greatest accuracy. If necessary, carefully dilute cultures prior to analysis so that all samples are approximately the same density. In field samples, increase the threshold, if possible, to eliminate noise without losing cells. To verify the efficiency of counting, prepare a series of k n o w n concentrations of beads or cells. Acquire counts for each dilution by cytometry and by an independent measure, such as microscopy or particle counter. Plot the cytometry counts vs. the expected counts to determine how the coincidence rate affects the accuracy of acquisition by the flow cytometer.
Data storage/archival Careful consideration of data archival is necessary owing to the large size of m a n y listmode files. An active laboratory can produce several gigabytes of data quickly, so an adequate capability for backup and archival is essential. Currently, the most convenient and cost-effective method for data archival is on compact discs (CDs). Given the availability and recent reductions in cost of CD r e a d / w r i t e drives (CD-RW), CDs appear to be a better long-term storage m e d i u m than tape, which must be refreshed at annual intervals. Also, tape drives m a y fail, become outdated or unavailable, and m a y be incompatible with newer computers. CDs can be read from most platforms.
ee.e, ee, e, D I S C U S S I O N
AND
FUTURE
DIRECTIONS
W h a t we have learned about marine picoplankton from flow cytometry In 1989, an entire issue of Cytometry was devoted to cytometry in the aquatic sciences in recognition of the potential of this powerful technology in oceanography. The areas highlighted included both ecological questions and optical properties. The ability to examine heterogeneity in size, abundance and activities of cells was exciting. Although a majority of the studies related to phytoplankton, the innovative studies of Button and Robertson (1989) showed that bacterial biomass could be determined from flow cytometric measurements and then used to improve estimates of bacterial activity measurements. In the ensuing years, the importance of biomass and activity measurements continues to be a major area of research. The importance of heterogeneity in cellular properties has been reinforced (Davey and Kell, 1996). The discovery of Prochlorococcus by Chisholm and co-workers (1988) was one of the most significant accomplishments in the past decade, and m a d e possible only because flow cytometric analyses were conducted at sea. The recognition that Prochlorococcus is probably the most abundant photosynthetic organism in the ocean has had a tremendous impact on our understanding of the
333
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contribution of picoplankton to the food web of the ocean. Consequently, in studies of community structure throughout the world's oceans, flow cytometry is now an essential tool (Li and Wood, 1988; Olson et al., 1990; Landry et al., 1995a; Binder et al., 1996; Campbell et al., 1997; 1998; Gin et al., 1999; Tarran et al., 1999). Improvements in our protocols for examination of bacterial populations has led to the extension of this technology to virus populations in the ocean (Marie et al., 1999a). Along with the remarkable discoveries afforded by pushing this technology to its limit, other significant applications have resulted from using the technology at its best: analyzing thousands of cells. Thus, in processoriented studies, enumeration by flow cytometry has improved our capabilities as well. Examples include applications of the cell cycle method for determination of species-specific growth rates (Carpenter and Chang, 1988) to microbial populations (Vaulot et al., 1995; Liu et al., 1997, 1998; Shalapyonok et al., 2000), grazing studies and trophic dynamics (Cucci et al., 1989; Landry et al., 1995 a,b; 1998). The number of applications continue to grow, and flow cytometry is now successfully utilized in many studies of bacterial community structure, composition and activity (Fouchet et al., 1993; Collier and Campbell, 1999; Servais et al., 1999).
Advantages and limitations One of the advantages of flow cytometry is that tile rapid analysis of a large number of events (10 000 or more) provides results with a greater level of statistical significance than traditional or epifluorescence microscopy (in which only several hundreds of cells are enumerated). The limitation of flow cytometry, however, is that taxonomic resolution for larger cells is not possible with standard parameters (with the exception of the high scatter or time of flight parameters for pennate diatoms, Olson et al., 1991). Thus, the overall resolution may be less than the class-level partitioning of autotrophic biomass possible with HPLC. However, recent studies have provided estimates of total biomass from size spectra and abundance data obtained by flow cytometry and carbon conversion factors specific for each component (Shalapyonok et al., 2000; Garrison et al., 2000). Cell volume, as well as biomass, has also been estimated from FALS signals of extremely small marine bacteria (Koch et al., 1996). Better estimates of biomass and cell volume will enhance the value of flow cytometric data. Another advantage of flow cytometry is that routine operation at sea is possible; consequently, real-time analyses are possible for help in designing field experiments, as well as conducting diel experiments. Both the EPICS 753 (Olson et al., 1991) and the FACSCalibur (Li, 1989; 1994) have been used extensively at sea. Currently, the modern benchtop flow cytometers are the most cost-effective and the most compact for microbial identification and enumeration at sea. One limitation of this instrument, however, is that it may not be sensitive enough to detect all surface populations of Prochlorococcus. This is especially true in surface samples from highly oligotrophic waters where cellular fluorescence is extremely weak; thus, Prochlorococcus populations can be 'lost' into the background noise
334
of the system. In such cases, samples can be analyzed on an EPICS 753, operating with all lines and at 4 W to obtain sufficient power to see Prochlorococcus in surface samples (Olson et al., 1990; Campbell et al., 1997). As an alternative, Frankel and co-workers (1990) modified a FACSCan instrument to increase its sensitivity for analysis of picoplankton. For most standard equipment, however, this remains a limitation. For the greatest success, it is best to examine live samples and thereby avoid artefacts of fixation (e.g. glutaraldehyde-induced autofluorescence, cell loss). When live sampling is not possible, fixation and freezing have proved effective for many, but not all cell types (Vaulot et al., 1989). Hence, many larger, fragile eukaryotic algae may not be adequately represented in preserved samples. Fortunately, most prokaryotic cells withstand this protocol better than the more fragile phytoplankton species. The constant supply of new stains and fluorescent probes for functional activity have provided a considerable array of new tools. Most must be optimized for marine applications since properties in seawater are likely to be very different than in the mammalian cells for which they were designed. Optimal performance of most flow cytometers depends on the population abundance. The cells of interest should be present at a level of at least 10~ml '; thus, flow cytometry is appropriate for most picoplankton populations. For cells present at abundances less than 10~ml ~,most methods of concentration (tangential flow, centrifugation) have proven unsatisfactory for quantitative recovery, so methods to increase the sample volume delivered are preferred (Olson et al., 1989; Peeters et al., 1989; Dubelaar and van der Reijden, 1995; Cavendar-Bares et al., 1998). In working with the larger members of the microbial community, clogging of the flow cell, or orofice, is a potential problem. In coastal samples where larger cells or aggregates of cells may be present, pre-filtration is recommended. Samples should be drawn through Nitex screening (e.g. 53 l~m Nitex when using a 76 lam jetin-air nozzle (Olson et al., 1993) or 35 pm Nitex-capped tubes (Falcon). The ability to physically separate a specific population from a sample is one of the major attractions of flow cytometric analysis. Specific protocols for sorting will depend on the instrument, but if a population can be distinguished by the standard analyses discussed here, they should then be readily sorted. With the FACSCalibur instrument, sorting is very simple to set up, however, sorting is accomplished using a mechanical sliding mechanism and thus is limited to 300 particles s ', which is considerably slower than other instruments and so may be limiting in some cases. For applications in which microbial populations are sorted additional precautions, in particular sterility, must be considered (Harkins, 1999). In addition, the rate of recovery is never 100(g. Due to sort logic and coincidence, some cells are rejected, so it is always necessary to verify abundances in sorted samples if quantification analysis is the objective.
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Studies of marine microbial ecology will continue to benefit from advances in the techniques and instrumentation developed for flow cytometry. Future applications will certainly include more specific probes
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with which to identify and quantify specific cell types and processes (Wallner et al., 1997; Sieracki et al., 1999). Thus, the promise of better taxonomic approaches awaits us, as does the possibility for a better understanding of the activity of individual microbial populations. New modifications to instruments will no doubt also provide further opportunities to examine processes and properties at the single-cell level. For example, photosynthetic parameters determined from active fluorescence measurements have been measured using a recently designed flow cytometer (Olson et al., 1999). Such 'pump-during-probe' measurements on individual cells provide a measure of heterogeneity among phytoplankton cells and contribution of individual cell types to bulk signals. Improved sorting technology, for example the MoFlo (Cytomation, Inc., Ft. Collins, CO) and HyPerSort (Coulter) systems, will provide improved capabilities for isolating populations for subsequent analyses. In many systems, the charged droplets are sorted based on two parameters: positive or negative charging, thus limiting the sort to two populations. The new MoFlo cytometer can sort four samples at a time because the events are charged within a range of electrical charges. There have been several examples of sorting in combination with field experiments (Li, 1994; Lipshultz 1995; Moore et al., 1998; Urbach and Chisholm, 1998), and with improved capabilities no doubt there will be many more. Flow cytometry has provided microbial ecologists the capability to identify and quantify the picoplankton more rapidly and with improved precision over traditional microscopy. This is especially true for Prochlorococcus, for which there is no reliable alternative. In providing a single-cell approach, flow cytometry permits questions to be addressed on a scale that is more relevant to microbial populations than bulk measurements. Finally, the tremendous potential of molecular probes in combination with flow cytometry will further enhance our understanding of the dynamics and diversity of the photosynthetic picoplankton.
RESOURCES The flow cytometry community spans quite a number of biological disciplines. The International Society for Analytical Cytometry (ISAC) publishes the journal, Cytometry, and holds annual international meetings. ISAC Office
60 Revere Drive, Suite 500 Northbrook, IL 60062-1577, USA Phone: 847-205-4722 Fax: 847-480-9282 e-mail: [email protected] http://www.isac-net.org/
Cytometry http://www.interscience.wiley, com/jpages / 0196-4763 / 336
There are also a n u m b e r of World Wide Web pages maintained by some of the major flow c y t o m e t r y laboratories. Both the Scripps Research Institute and the Salk Institute maintain extensive lists of information on flow c y t o m e t r y protocols, lasers, reagents, and useful software. The P u r d u e University is particularly helpful and dedicated to educational efforts. Many useful sites are linked from these pages, which should be consulted to obtain URLs for additional flow c y t o m e t r y sites. Salk Institute Flow C y t o m e t r y Laboratory h t t p : / / p i n g u . s a l k . e d u / fcm.html The Scripps Research Institute http://facs.scripps.edu/ P u r d u e University http://flowcyt.cyto.purdue.edu/index.htm To join an email discussion group: http://flowcyt.cyto.purdue.edu/hmarchiv/index.htm Aquatic flow c y t o m e t r y mailing list http://www.flowcytometry.org/
Acknowledgments I acknowledge funds from Texas A&M University and a research grant from the NSF in support of the acquisition of the flow cytometer at Texas A&M. I also thank the Texas A&M Faculty d e v e l o p m e n t p r o g r a m for providing funds to s u p p o r t this research and the preparation of this manuscript.
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Frankel, D. S., Frankel, S. L., Binder, B. J. and Vogt, R. E (1996). Application of neural networks to flow cytometry data analysis and real-time cell classification. Cytometry 23, 290-302. Garrison, D. L., Gowing, M. M., Hughes, M. P., Campbell, L., Caron, D. A., Dennett, M. R., Shalapyonok, A., Olson, R. J., Landry, M. R., Brown, S. L., Liu, H., Azam, E, Steward, G. E, Ducklow, H. W. and Smith, D. C. (2000). Microbial food web structure in the Arabian Sea: A US JGOFS study. Deep-Sea Res. II. 47, 1387-1422. Gin, K. Y. H., Chisholm, S. W. and Olson, R. J. (1999). Seasonal and depth variation in microbial size spectra at the Bermuda Atlantic time series station. Deep-Sea Res. 46, 1221-1245. Harkins, K. R. (1999). Sorting of bacteria. In: Current Protocols in Cytometry (J. P. Robinson, Ed,), pp. 11.4.1-11.4.12. John Wiley and Sons, Inc., New York. Jeffrey, S. W., Mantoura, R. E C. and Wright, S. W. (eds). (1997). Phytoplankton Pigments i~ Oceanography. Mono~raphs on Oceanographic Methodology. UNESCO Publishing, Paris. Koch, A. L., Robertson, B. R. and Button, D. K. (1996). Deduction of the cell volume and mass from forward scatter intensity of bacteria analyzed by flow cytometry. J. Miclvbiol. Methods 27, 49-61. Landry, M. R., Constantinou, J. and Kirshtein, J. (1995a). Microzooplankton grazing in the central equatorial Pacific during February and August, 1992. Deep Sea Res. 42, 657-671. Landry, M. R., Kirshtein, J. and Constantinou, J. (1995b). A refined dilution technique for measuring the community grazing impact of microzooplankton, with experimental tests in the central equatorial Pacific. Mar. Ecol. Pro g. Set. 120, 53-63. Landry, M. R., Brown, S. L., Campbell, L., Constantinou, J. and Liu, H. (1998). Spatial patterns in phytoplankton growth and microzooplankton grazing in the Arabian Sea during monsoon forcing. Deep-Sea Res. 11 45, 2353 2368. Li, W. W. K. (1989). Shipboard analytical flow cytometry of oceanic ultraphytoplankton. Cytomett?/10, 564-579. Li, W. K. W. (1994). Primary production of prochlorophytes, cyanobacteria, and eucaryotic ultraphytoplankton: Measurements from flow cytometric sorting. Limuol. Oceanogr. 39, 169-175. Li, W. K. W. and Wood, A. M. (1988). Vertical distribution of North Atlantic ultraphytoplankton: analysis by flow cytometry and epifluorescence microscopy. Deep-Sea Res. 35, 1615-1638. Lipschultz, E (1995). Nitrogen-specific uptake rates of marine phytoplankton isolated from natural populations of particles by flow cytometry. Mar. Ecol. Prog. Scr. 123, 245-258. Liu, H., Campbell, L., Landry, M. R., Nolla, H. A., Brown, S. L. and Constantinou, J. (1998). Prochlorococcus and Synechococcus growth rates and relative contributions to production in the Arabian Seas during Southwest and Northeast Monsoons. Deep-Sea Res. 11 45, 2327-2352. Liu, H., Nolla, H. A. and Campbell, L. (1997). Prochlorococcus growth rate and contribution to primary production in the equatorial and subtropical North Pacific Ocean. Aquatic Microbiol Ecol. 12, 39-47. Marie, D., Partensky, E, Jacquet, S. and Vaulot, D. (1997). Enumeration and cell cycle analysis of natural populations of marine picoplankton by flow cytometry using the nucleic acid stain SYBR green I. Appl. Environ. Microbiol. 63,
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Simon, N., Campbell, L., Ornolfsdottir, E., Groben, R., Guillou, L., Lange, M. and Medlin, L. K. (2000). Oligonucleotide probes for the identification of three algal groups by dot blot and fluorescent whole-cell hybridization. J. Eukaryotic Microbiol. 47, 76-84. Stairs, J. R. M., Breedveld, L. W., Derksen, M. W. J., Kateman, G., Balfoort, H. W., Snoek, J. and Hofstraat, J. W. (1992). Pattern-classification with artificial neural networks--classification of algae, based upon flow cytometer data. Anal. Chem. 258, 11-25. Tarran, G. A., Burkill, P. H., Edwards, E. S. and Woodward, E. M. S. (1999). Phytoplankton comunity structure in the Arabian Sea during and after the SW monsoon, 1994. Deep-Sea Res. II. 46, 655-676. Troussellier, M., Courties, C., Lebaron, P. and Servais, P. (1999). Flow cytometric discrimination of bacterial populations in seawater based on SYTO 13 staining of nucleic acids. FEMS Microbiol. Ecol. 29, 319-330. Urbach, E. and Chisholm, S. W. (1998). Genetic diversity in Prochlorococcus populations flow cytometrically sorted from the Sargasso Sea and Gulf Stream. Limnol. Oceano~r. 43, 1615-1630. Vaulot, D. (1989). CYTOPC: processing software for flow cytometric data. Sigtlal alut Noise 2, 8. Vaulot, D., Marie, D., Olson, R. J. and Chisholm, S. W. (1995). Growth of Prochlorococcus, a photosynthetic prokaryote, in the equatorial Pacific Ocean. Scielzce 268, 1480-1482. Vaulot, D., Courties, C. and Partensky, E (1989). A simple method to preserve oceanic phytoplankton for flow cytometric analyses. Cytometry 10, 629-635. Veldhuis, M. J. W., Cucci, T. L. and Sieracki, M. E. (1997). Cellular DNA content of marine phytoplankton using two new fluorochromes: Taxonomic and ecological implications. J. Phycol. 33, 527-541. Vrieling, E. G. and Anderson, D. M. (1996). Immunofluorescence in phytoplankton research: applications and potential. J. Phycol. 32, 1-16. Wallner, G., Fuchs, B., Spring, S., Beisker, W. and Amann, R. (1997). Flow sorting of microorganisms for molecular analysis. Appl. Environ. Microbiol. 63, 4223-4231. Watson, J. V. (ed.) (1992). Flow Cytometry Data Analysis. Basic Collcepts and Statistics. Cambridge University Press, Cambridge. Weitzorrek, J., Plesnila, N., Baethmann, A. and Kachel, V. (1999). A new multiparameter flow cytometer: optical and electrical cell analysis in combination with video microscopy in flow. Cytontotry 35, 291-301. Wilkins, M. E, Morris, C. W. ~nd Boddy, L. (1994). A comparison of radial basis function and back propagation neural networks for identification of marinephytoplankton from multivariate flow-cytometry data. Comput. Appl. Biosci. 10, 285-294. Wood, J. C. S. (1997). Establishing and maintaining system linearity. In: CurreHt l)roh~cols in Cytometry (P. J. Robinson et al., Eds), pp. 1.4.1-1.4.12. John Wiley and Sons, Inc., New York. Yentsch, C. M., Horan, P. K., Muirhead, K., Dortch, Q., Haugen, E. M., Legendre, L., Murphy, L. S., Phinney, D., Pomponi, S. A., Spinrad, R. W., Wood, A. M., Yentsch, C. S. and Zahurenec, B. J. (1983). Flow cytometry and sorting: a powerful technique with potential applications in aquatic sciences. Limnol. Ocealloy, r. 28, 1275-1280.
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List of suppliers Amersham Pharmacia Biotech Ltd. Amersham Pharmacia Biotech AB SE-751 84 Uppsala, Sweden Tel: +46 (0) 18 612 O0 O0 Fax: 46 (0) 18 612 12 O0 800 Centennial Avenue P.O. Box 1327 Piscataway, NJ 08855-1327 USA Tel: 1-732-457-8000 Fax: 1-732-457-0557 http://www.apbiotech.com/
Cytomation, Inc. 4850 Innovation Drive Fort Collins, CO 80525 USA Teh 800-822-9902 or 970-226-2200 Fax: 970-226-0107 e-maih [email protected] h t tp :/ /www.cyt oma t ion .com
Fluorophores
DAKO DAKO Corporation 6392 Via Real Carpinteria, CA 93013, USA Teh 805 566 6655 Fax: 805-566-6688 http://www.dakousa.com h t tp :/ /www.dako.com /
Beckman Coulter, Inc. 4300 N. Harbor Boulevard, P.O. Box 3100 Fullerton, CA 92834-3100, USA Phone: 800- 233-4685 or 714-871-4848 Fax: 714-773-8283 h t tp :/ /www.beckman.com / source book: http://www.bdfacs.com/source_book/
Instruments ( EPICS@ ALTRATM and HyPerSortTM, Coulter@ EPICS@ XL); calibration beads Becton Dickinson BD hnmunocytometry Systems 2350 Qume Drive, San Jose, CA 95131-1807, USA Tel: 800-223-8226 Customer Support: 800-448-2347 Fax: 408-954-2347 http://www.bdfacs.com/
Instruments, calibration beads, immunochemicals Bio-Rad Laboratories, Inc. Diagnostics Group 4000 Alfred Nobel Drive Hercules, CA 94547, USA Phone: 800-224-6723 http://www.bio-rad.com
Instruments, software
Immunochemicals DeNovo Software 64 McClintock Crescent Thornhill, Ontario L4J 2T1, Canada Tel: +001-905-738-9442 Fax: +001-905-738-5126 [email protected] h ttp://www.denovosoftware.com/
Analysis software Jackson |mmunoResearch Laboratories, Inc. P.O. Box 9 872 West Baltimore Pike West Grove, PA 19390, USA Teh 800-367-5296 or 610-869-4024 Fax: 610-869-0171 e-maih [email protected] http://www.jacksonimmuno.com/
Immunochemicals
Instruments (BRYTE HS)
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Molecular Probes, Inc. PO Box 22010 Eugene, OR 97402-0469, USA 4849 Pitchford Ave. Eugene, OR 97402-9165, USA Td: (541) 465-8300 Fax: (541) 344-6504 http://www.probes.com/ catalog: http://www.probes.com/handbook/ sections/OOOO.html Stains; fluorophores; calibration standards Omega Optical, Inc. P.O. Box 573 Brattleboro, VT 05302-0573, USA Teh 802-254-2690 Fax: 802-254-393 7 e-maih [email protected] http: //www.omegafilters.com Optical filters Partec GmbH Otto-Hahn-Str. 32 D-48161 Miinster, Germany Teh 49-2534-8008-0 Fax: 49-2534-8008-90
PolySciences 400 Valley Road Warriny, ton, PA 18976, USA 215 343-6484 800 523-2575 Fax: 800-343-3291 http://www.polysciences.com/ Calibration standards SpheroTech, Inc. 1840 industrial Dr. Suite 270 Libertyville, IL 60048-9817, USA 800-368-0822 847-680-8927 [email protected] http://www.sphetvtech.com Calibration standards Verity Software House, Inc. PO Box 247 45A Augusta Road Topsham, ME 04086, USA Teh 207-729-6767 ext.190 Fax: 207-729-5443 e-mail [email protected] http://www.vsh.com/ Analysis software
Instrument (particle analysing system) Phoenix Flow Systems, Inc. 11575 Sorrento Valley Road #208 San Diego, CA 92121, USA Teh 800-886-FLOW (3569) (619) 453-5095 Fax: (619) 259-5268 e-maih [email protected] Analysis software
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17 Collection and Identification of Marine Yeasts
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Jack W Fell
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CONTENTS Introduction Methods of collection Methods of isolation Methods of species identification Methods of strain identification Future directions
~,~,~,~,4,~, I N T R O D U C T I O N Yeasts are a polyphyletic group of basidiomycetous and ascomycetous fungi with a unifying characteristic of a unicellular growth stage. There are approximately 100 genera and 800 described species of yeasts (Kurtzman and Fell, 1998); estimates suggest that these numbers represent l% of the species that exist in nature. Their environmental role is similar to many other fungi, acting as saprophytes by converting plant and animal organics to yeast biomass and to by-products, which may have commercial importance. Some yeasts are pathogenic to plants and animals. Yeasts are found throughout aerobic marine habitats; their numbers and species distributions are dependent on concentrations and types of organic materials. Nearshore environments are usually inhabited by tens to thousands of cells per liter of water, whereas low organic surface to deep sea oceanic regions contain 10 or fewer cells 1 ', although local nutrient pulses may foster concentrations of yeast cells that reach 3-4 thousand 1 '. These numbers reflect yeasts that can be cultured on artificial growth media; the potential numbers of unculturable yeasts in marine environments, or any other habitat, is unknown. This chapter discusses methods of isolation and collection of yeasts using traditional techniques, and identification of species and strains with molecular methods. METHODS IN MICROBIOLOGY, VOLUME 30 ISBN ()~12-521530 4
Copyright © 2001 Academic Press Ltd All rights of reproduction in anv form reserved
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METHODS
OF COLLECTION
Techniques for collecting yeasts should be tailored to the environment. The majority of yeasts, and other fungi, are obligate aerobes that require oxygen for growth and reproduction. Therefore, yeasts usually do not inhabit anaerobic waters and sediments, although they can be preserved under special conditions such as deep layers of ice. Water and sediment samples from shallow water environments can be manually collected using sterile jars, vials and plastic bags. Deep water sediments can be remotely collected using traditional grabs, gravity and piston cores or submersibles. Deep water samples can be collected using Sterile Bag Samplers (General Oceanics Model 10301.5, web site www.generaloceanics.com). Nearshore water samples of 250 ml are generally adequate, whereas open ocean samples of I 1 or more are routinely required. Sediment samples in volumes of approximately 50ml are sufficient. Sample design should recognize the patchy distribution of populations to include adequate replicate samples.
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METHODS
OF I S O L A T I O N
Traditional methods of yeast isolation and cultivation, which are successfully employed for obtaining large numbers of isolates, have specific limitations. The culture media and environmental growth conditions (particularly temperature) are selective; rapid growing strains will overgrow slower growing species; and plate techniques are limited to a maximum number of cells (-300) that can be accommodated on a growth plate, consequently r a r e species may not be represented. Additionally, cell numbers obtained with plate cultivation techniques do not reflect factors such as turnover rates, hyphal fragmentation, spore release or rates of consumption by various invertebrates. To overcome these limitations, careful consideration of environmental conditions is necessary. A variety of media and incubation conditions can be employed and designed by the researcher. The method that we use for water sampling employs filtration through 47 mm diameter nitrocellulose filters 0.45 t2m pore-size, using an autoclavable glass or plastic filter apparatus (Millipore Corp). The filter is placed face-up on a nutrient agar medium. A variety of general and specific media have been described (Yarrow, 1998). A widely used medium is Wickerham's YM medium that contains 0.3% yeast extract, 0.3% malt extract, 0.5% peptone, 1% glucose and 2% agar prepared with seawater at a salinity equivalent to the sample site. Bacteria are inhibited by addition of chloramphenicol (200 mg 1 ~) to the medium prior to autoclaving. An alternative is an antibiotic mixture of penicillin G and streptomycin sulfate (each at 150-500 mg 11), added dry to autoclaved, cooled (45°C) medium. Sediment particles can be placed directly on an agar medium or known quantities of sediment can be placed in a test tube with a given volume of sterile seawater, vortexed and diluted 1:10 in a sterile seawater series, 348
followed by preparation of standard spread plates from each of the dilution series. Growth plates are incubated at temperatures designed to maintain ambient environmental conditions. For example, polar and deep-sea samples should be incubated at ~5°C. Temperatures required for temperate and tropical samples often result in overgrowth by filamentous fungi, which can be reduced by incubation at temperatures ~12°C. Growth plates should be inspected daily. Suspected yeast colonies can be picked and transferred to a microscope slide for inspection. Confirmed yeast growth can be transferred from the isolation medium to a growth medium (YM seawater agar lacking antibiotics). Streak cultures should be prepared to ensure purity of the isolate. Representative colonies can be transferred to a slant culture tube for further study.
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METHODS
O F SPECIES I D E N T I F I C A T I O N
Classical techniques Classical techniques for the identification of species of yeasts require morphological, physiological and biochemical tests. These methods have been presented in detail by Yarrow (1998). Generally, these techniques take 2-3 weeks to complete and the results may be ambiguous. Consequently, molecular methods, which will be described below, have been devised and are becoming routine in some laboratories. Species can be identified by sequence analysis of purified isolates or via species-specific primers and hybridization probes from purified isolates or from field samples of mixed species.
Molecular sequence analysis Research on ascomycetous (Kurtzman and Robnet, 1998) and basidiomycetous yeasts (Fell et al., 2000) resulted in a D1/D2 region large subunit ribosomal DNA (LrDNA) data bank of all described species, which is accessible in GenBank. A second data bank of the ITS region is under development. Identification of a yeast isolate takes the following steps: (1) DNA extraction from a fresh culture; (2) amplification of the - 2 kb rDNA fragment from primer NS7 to LR6 (Figure 17.1); (3) sequencing of D1/D2 a n d / o r ITS regions, (4) sequence analysis and species identification. D N A extraction. There are numerous methods for DNA extraction and purification. The method followed in our laboratory is a modified QIAamp DNAeasy tissue culture procedure (Qiagen, Inc. Cat. # 69506). Yeast and other fungal cell walls are often difficult to break to extract DNA. Therefore, we use a lysing enzyme incubation to prepare protoplasts, from which the DNA is extracted and purified.
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Step 1. Cells from pure cultures are grown for 12-14 h in GYP broth (2% glucose, 0.5% peptone and 0.1% yeast extract). Prepare protoplasts as follows: 1. Spin cells at top speed in a microfuge for 5 min. 2. Decant supernatant and add 1 ml of sterile water, vortex to release pellet. 3. Spin again at top speed for 5 rain. 4. Decant supernatant and dissolve (vortex) pellet in 1 ml of 20 mM sodium citrate, pH 5.8/1 M sorbitol, containing 1 0 m g m l ~ Lysing Enzyme from Trichoderma harzianum (Sigma Cat. # L1412) (freshly prepared for each procedure). 5. Incubate for 2 h at 37°C. 6. Spin at 5000 rpm for 2 min, decant supernatant. Follow standard DNAeasy protocol as follows: 7. Add 180 pl of buffer ATL, vortex to release pellet (if ATL has precipitated, put bottle in waterbath). 8. Incubate at 55°C for 30 min to get DNA into solution, vortex and check that pellet has dissolved. 9. Add 200 1.11of buffer AL, vortex, incubate at 70°C for 10 min. 10. Add 210 lal of 95% ETOH, vortex. 11. Apply lysate to spin column, spin for 1 min (full speed). 12. Discard filtrate and wash by adding 500 tJl of buffer AWl, spin for 1 min, discard filtrate. 350
13. Wash with 500 1-llof buffer AW2, spin for 3 min, discard filtrate. 14. Place column in a 1.5 ml eppendorf tube. 15. Warm 150 Ill of buffer AE in a separate tube to 70°C. (Add less volume to concentrate DNA.) 16. Add warmed buffer to column, let it stand for 1 min and spin for 1 min. The resulting DNA solution can be amplified. Step 2. Amplify ribosomal DNA fragment using standard thermal cycle protocol and primers NS7 and LR6 (Figure 17.1, Table 17.1). Various enzymes are available; we use DYNAzyme EXT DNA polymerase (MJ Research Cat#F505s). Amplicons should be purified and a variety of techniques are available. We use the QIAquick PCR Purification Kit (Qiagen Cat. # 28106) following the Qiagen protocol.
Table 17.1 Primers for amplification and sequencing of yeast r D N A (see Figure 17. I ) Primer
Sequence
NS7 1TS5 ITS1 ITS3 F63 1TS4 R635 LR6 LR11F LR12F IGIF 5SF 5SR NS1R SR3R SR1R
GAG GCA ATA ACA GGT CTG TGA TGC GGA AGT AAA AGT CGT AAC AAG G TCC GTA GGT GAA CCT GCG G GCA TCG ATG AAG AAC GCA GC GCA TAT CAA TAA GCG GAG GAA AAG TCC TCC GCT TAT TGA TAT GC GGT CCG TGT TTC AAG ACG G CGC CAG TTC TGC TTA CC TTA CCA CAG GGA TTA CTG GC CTG AAC GCC TCT AAG TCA GAA CAG ACG ACT TGA ATG GGA ACG GCA CCC TGC CCC GTC CGA TCC GGA TCG GAC GGG GCA GGG TGC GAG ACA AGC ATA TGA CTA C GAA AGT TGA TGA GGC T ATT ACC GCG GCT GCT
Step 3. DNA can be sequenced with standard protocols of any of the commercially available automated DNA sequencers. Our laboratory uses the LiCor NEN Global IRe DNA Sequencer. Sequencing primers for the D1/D2 and ITS regions (Figure 17.1) are listed in Table 17.1. Forward and reverse strands should be sequenced, compared and the final sequence corrected. Step 4. The acquired D1/D2 sequence can be compared against available sequences in GenBank using a BLAST search at the NIH web site:
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http://www.ncbi.nlm.nih.gov/BLAST/. That search will provide a pairwise alignment of species with identical and similar sequences. If the sequence is identical to a known species, the biology of the species and other background information is available in Kurtzman and Fell (1998). If the GenBank match relates to a strain with a Centraalbureau voor Schimmelculture (CBS) accession number, the physiological test and historical data can be obtained at the CBS web site: http://www.cbs.knaw.nl/www/search ydb.html. Ascomycete data (Kurtzman and Robnet, 1998) was entered in GenBank with NRRL Yaccession numbers. Cross-reference of Y and CBS numbers can be obtained at the USDA web site: http://nrrl.ncaur.usda.gov/; the physiological data is not available at the USDA site. If there is a question regarding the relationship of an isolate to a GenBank-identified species, confirmation or rejection of the identity can be determined by the additional sequence analysis of the ITS region. Data accumulating in our laboratory demonstrates that differentiation of closely related species requires analysis of both D1/D2 and ITS regions. Phylogenetic positions of ascomycetous species can be determined by examination of the trees presented by Kurtzman and Robnet (1998), basidiomycete trees are available in Fell et al. (2000). As an example, a strain identified as the marine-occurring species Cystofilobasidium bisporidii, would belong to the Cystofilobasidiales of the Hymenomycetes as demonstrated in the example (Figure 17.2) of a D1/D2 large sub-unit rDNA phylogenetic tree. The phylogenetic position of isolates, which differ from all known species in GenBank, can be estimated by a comparison of results of the GenBank BLAST search to the phylogenetic trees. Sequences of related species can be imported from GenBank into an alignment program, such as ClustalX, which is available at http://www.icgeb.trieste.it/net/ clustalx/clustalx.html. The alignment results, coupled with the program treeview (http://taxonomy.zoology.gla.ac.uk/rod/treeview.html), provide a preliminary phylogenetic analysis, which can be further analyzed with PAUP* 4.0 (Sinauer Press: www.sinauer.com/Titles / frswofford.htm). Another useful program, Lasergene99 (http:// www.dnastar.com/), combines the features of Clustal and treeview that can be integrated with a BLAST search.
PCR primers Species identifications with sequence analysis, although accurate, are time consuming and expensive when the goal may be to survey for specific species. Species-specific PCR primers can be designed by examination of sequence alignments of species clusters as depicted in Figure 17.2. Through the use of an alignment program such as ClustalX, species specific regions can be detected. Primers are usually designed at 18-20 nucleotides in length and differ from other species sequences by, at least, two nucleotides at the 3' end of the primer. Generally, the 3' terminal primer should be a G or C. Specificity of the primer can be determined by
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Figure 17.2. Example of a yeast phylogenetic tree: hymenomycetous yeasts: phylogenetic analysis (PAUP 4.0) of the D1/D2 region of the large subunit rDNA. One of 100 equally parsimonious trees. Number of characters - 651, constant characters = 285, parsimony-uninformative characters - 71, parsimony-informative characters = 295. Tree length 2163, consistency index (CI) - 0.2779, retention index (RI) = 0.7765.
353
a BLAST search. Characteristics of the primer (melting temperature, percent GC, presence of secondary structure and primer dimers) can be determined by the Sigma-Genosys calculator (http://www. genosys.com/cgi-win/oligo calconly.exe). The most useful primers will be at least 200 nucleotide positions from either of the universal primers in order to produce an amplicon that is easily distinguished from a universal amplicon. The following method is a modified three-primer technique (Fell, 1993, 1995), using two universal primers and a species-specific primer that is interior in the amplicon to the universal primers. The three primers are used in equal concentrations in a single PCR reaction with the test DNA. If target DNA is present, the species-specific amplicon is detected, whereas non-target DNA results in the universal amplicon.
Method
Target DNA. DNA is extracted as above, however, the isolates do not have to be purified by streak plate isolations. The technique is applicable to the identification of species within mixed populations. As an example of an identification using the D1/D2 region, master mixes can be prepared, as follows, in multiples using DYNAzyme EXT DNA polymerase protocol. 1 pl DNTP 5 pl 10x DNAzyme (511) buffer 1 pl DYNAzyme EXT DNA polymerase 5 pl genomic DNA 1 121LR6 primer (25 pM) 1 1_11F63 primer (25 pM) 1 pl species specific primer (25 pM) 35 121water Amplification (using an MJ Research PTC 100) with the following reference program: 94°C for I min followed by 30 cycles of 94°C for i min, 58°C for 1 min and 72°C for 1 min with a final extension at 70°C for 8 min. However, the program will require experimentation with annealing temperature based on the melting temperatures of the primers. Results can be visualized with 1.0% agarose TPE (0.09 M Tris-phosphate, 0.002 M EDTA) gels stained with ethidium bromide. In the presence of the species-specific target DNA, the smaller species-specific amplicon and usually the (-1200 bp) universal amplicon will be present. In the absence of the species-specific target DNA, the universal amplicon is present. Other polymerases can be used (Fell, 1995), such as Tth DNA polymerase (Promega). However, Taq DNA polymerase does not provide species-specific amplicon results with the three-primer technique. Taq is better suited to a two-primer technique using a speciesspecific primer and a single universal primer (Mannarelli and Kurtzman, 1998).
354
~ . ~
M E T H O D S OF S T R A I N I D E N T I F I C A T I O N e,,
Differentiation of strains within a species can play a significant role in ecological population analyses. The intergenic spacer region (IGS) has been studied for population and strain level differences among a wide range of eukaryotes, including plants (Cordesse et al., 1993), invertebrates (Crease, 1995) and yeasts (Fell and Blatt, 1999; Diaz and Fell, 2000). The IGS region is between the large and small rDNA genes and includes the 5S gene. There is considerable IGS variability within and between species. For example, differentiation of strains of Phaffia was determined by indels among a series of sub-repeats (Fell and Blatt, 1999), whereas Mrakia strains were determined by nucleotide substitutions (Diaz and Fell, 2000). Due to the variability between species, considerable experimentation must be undertaken with each species to ascertain amplifying and sequencing primers. Table 17.1 provides a list of useful primers.
FUTURE DIRECTIONS The PCR detection technique is useful for identification of small numbers of species, however, when several species need to be identified, the PCR matrix is difficult to handle. Therefore, more appropriate techniques will include hybridization probes with macro- and microarrays designed to identify large numbers of species. These hybridization arrays can be used to directly detect cells in water or solid substrates in the absence of growth techniques. Another significant step will be the development of inexpensive quantitative assays. All of these techniques, which are under development, required the initial production of a molecular database for species discriminations and hybridization probe design.
Acknowledgment The research clescribed and manuscript preparation were funded by the Ocean Sciences Division (Biological Oceanography) of the National Science Foundation.
References Cordesse, F., Cooke, R., Tremousaygue, D., Grellet, E and Delseny, M. (1993). Fine structure and evolution of the rDNA intergenic spacer in rice and other cereals. J. Mol. Evol. 36, 369-379. Crease, T. ]. (1995). Ribosomal DNA evolution at the population level: nucleotide variation ill intergenic spacer arrays of Daplmia puh:x. Genetics 141, 1327-1337. Diaz, M. R. and Fell, I. W. (2000). Systematics of psychrophilic yeasts in the genus Mrakia based on ITS and IGS rDNA sequence analysis. AHtonic va~z Lceuwenhoek 77, 7-12. Fell, J. (1993). Rapid identification of yeast species using three primers in a polymerase chain reaction. Mol. Mar. Biol. Bioteclmol. 3, 174-180.
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Fell, J. W. (1995). rDNA targeted oligonucleotide primers for the identification of pathogenic yeasts in a polymerase chain reaction. J. Ind. Microbiol. 14, 475-477. Fell, J. W. and Blatt, G. (1999). Separation of strains of the yeasts Xanthophyllomyces dendrorhous and Phaffia rhodozyma based on rDNA IGS and ITS sequence analysis. J. Ind. Microbiol. Biotech. 21, 677-681. Fell, J. W., Boekhout, T., Fonseca, A., Scorzetti, G. and Statzell-Tallman, A. (2000). Biodiversity and systematics of basidiomycetous yeasts as determined by large subunit rD1/D2 domain sequence analysis. Int. J. Syst. Evol. Microbiol. 50, 1351-1371. Kurtzman, C. P. and Fell, J. W. (1998). The Yeasts, A Taxonomic Study, 4th edn. Elsevier, Amsterdam. Kurtzman, C. P. and Robnett, C. J. (1998). Identification and phylogeny of ascomycetous yeasts from analysis of nuclear large subunit (6S) ribosomal DNA partial sequences. Antonie van Leeuwenhoek 73, 331-371. Mannarelli, B. M. and Kurtzman, C. P. (1998). Rapid identification of Candida albicans and other human pathogenic yeasts by using short oligonucleotides in a PCR. J. Clin. Microbiol. 36, 1634-41. Yarrow, D. (1998). Methods for the isolation, maintenance and identification of yeasts. In: The ~'asts, A Taxolzomic Study, 4th edn (C. P. Kurtzman and J. W. Fell, Eds), pp. 77-100. Elsevier, Amsterdam.
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18 Fungal Biomass and Productivity e.
S Y Newell Marine Institute, University of Georgia, Sapelo Island, Georgia, USA
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CONTENTS
Introduction Acetate-to-ergosterol (Ac-~ERG) method Conclusion Future directions
INTRODUCTION The Fungi constitute a Kingdom of evolutionarily closely related microbes that evolved along the same line leading away from the ancestral heterotrophic flagellate as did the animals and choanoflagellates (Kendrick, 1992; H a w k s w o r t h et al., 1995; Alexopoulos et al., 1996; DeLong, 1998). Most species in the Kingdom Fungi are mycelial (but see Chapter 17) eukaryotic organoosmotrophs with non-motile propagules (e.g. ascospores [sexual] and conidia [asexual]) and haploid status for most of the life cycle. The Chytridiomycota, believed to be near the root of the fungal evolutionary tree (Hawksworth et al., 1995), are exceptional in that their propagules are zoospores, and they are not mycelial (though some are rhizomycelial). There is a morphologically strikingly fungal-like group of zoosporic eukaryotic mycelial organoosmotrophs, the Oomycota, which were formerly considered to be primitive fungi, but are now known to be more closely related to diatoms than to fungi (Newell, 1996a; Delong, 1998; Fell and Newell, 1998; Dick et al., 1999), and to be members of the Kingdom Straminipila (Beakes, 1998). Thus, the oomvcotes evolved their mycelial subtrate-pervasion strategy independently of the true fungi. Because fungi digest natural materials such as saltmarsh grass from within the opaque solid substrate (Newell et al., 1996), their masses are best measured via biochemical indices (Newell, 1992). Fungi (including chytrids) and oomycotes are part of the marine microbiota (Fell and Newell, 1998), but biochemical-index methods have been developed for biomass and productivity measurement with natural samples only for METHODS IN MICROBIOLOGY,VOLUME3{) ISBN (1-12 521530-4
Copyright O 20{}1AcademicPress Ltd All rights of reproduction in any form reser\ ed
true fungi (Newell, 2000; Gessner and Newell, 2001). Biomasses of zoosporic, non-mycelial true fungi (the chytrids) can be measured using hexosamine techniques, but not with the ergosterol technique that is the focus of this chapter, because chytrids do not synthesize ergosterol (Weete et al., 1989). The oomycotes are also incapable of synthesizing ergosterol, and most do not synthesize chitin, so they are currently beyond the reach of published biochemical-index methods of mycelialmicrobial biomass analysis (Newell, 1994a; Fell and Newell, 1998). Therefore, this chapter, with its sterol-analytical emphasis, is directed at the mycelial true fungi. Mycelial fungi have a negligible presence in the marine plankton (Newell, 1996a) (but see Chapter 17). Fungi have their greatest impact in ecotonal marine ecosystems (marine/terrestrial transitions), where they can target senescing or dead parts of vascular plants for decay (saltmarsh grasses, mangroves) (Newell, 1994a; 1996a; Fell and Newell, 1998). Ascomycetes are the predominant fungi in these types of marine ecosystems (Kohlmeyer and Volkmann-Kohlmeyer, 1991; Fell and Newell, 1998; Kohlmeyer et al., 1999), including those impacted by pollution (Newell and Wall, 1998). Ascomycetous saltmarsh fungi are capable of digesting all parts of saltmarsh grasses, including the lignocellulosic structural framework (Newell at al., 1996). Saltmarsh-fungal production is efficient and can occur at high rates throughout the range of smooth-cordgrass (Spartina alterniflonT) marshes (Newell and Porter, 2000; Newell et al., 2000). There is a clear fungal connection into the saltmarsh foodweb: at least three shredder invertebrates (e.g. saltmarsh periwinkles), which avoid the bitter (ferulic acid) living shoots, are adapted for grazing of the standing, fungal-decayed shoots (Newell and Porter, 2000; Graqa et a[., 2000). It is possible to measure fungal biomass via biochemical proxies in two very different ways: as total mass (living plus dead, evacuated hyphae) or as living mass (membrane-containing mycelium). The hexosamine methods yield total-mass values (since chitin of cell walls is not readily lysed upon hyphal death), and the ergosterol methods yield living-mass values (since ergosterol is a membrane component, and membranes are readily lysed upon hyphal death) (Ekblad et al., 1998; Newell, 2000; Gessner and Newell, 2001). The ergosterol method was first described by agriculturally oriented scientists (Seitz ef al., 1979), but marine ecologists followed closely behind with their own version of the method (Lee el al., 1980), and it was marine microbial ecologists who extended the method beyond biomass measurement to measurement of productivity (Newell and Fallon, 1991). Aquatic microbial ecologists have provided subsequent, invaluable papers on development of the ergosterol-based, fungal-productivity method (Suberkropp and Weyers, 1996; Gessner and Chauvet, 1997). In this chapter the method for finding living-fungal mass and fungal productivity is described; see Newell (2000) and Gessner and Newell (2001) for description and references for hexosamine methods.
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A C E T A T E - T O - E R G O S T E R O L (Ac--~ERG) METHOD
Background and principle "o
Virtually all ascomycetes have ergosterol as the primary membrane sterol (exceptionally brassicasterol may be present in greater quantity than ergosterol), and no plants serving as fungal substrates synthesize ergosterol (Newell, 1992; Gessner and Newell, 2001), so ergosterol is an effective molecular marker for fungal presence in decaying plant material. Ergosterol is also a definitive proxy for fungal mass: only a few microbes other than fungi (green algae and protozoa) have been found to synthesize ergosterol (Newell, 1992; 2000; Gessner and Newell, 2001), and these species are not likely to be firmly bound to decaying leaves or stems of vascular plants in large quantities. The content of ergosterol in ascomycetous mass is rather constant; a recent assay for five species of ascomycetes in two distinct higher taxa (two replicate strains each) and from two different saltmarsh-grass species gave a homogeneous mean ergosterol content of 6.2 l,lg rag' mycelial organic mass (CV = _+8~7,) (Newell, 1996b). This mean value was only 25% different from the suggested general average value of 5 mg ergosterol per g organic fungal mass (Gessner and Newell, 2001; see also Djajakirana et al., 1996). Thus, ergosterol values can be converted into fungal-mass values without the risk of large error due to variation in conversion factors. Ergosterol has basic characteristics that make it a very practicable fungal proxy. It absorbs ultraviolet (UV) light with a peak at 282 nm, a rare feature among sterols (Newell, 1992), reducing the possibility of interference during chromatography. The strong absorbance at 282 nm has permitted development of very sensitive methods for ergosterol assay; e.g. the microwave-extraction method of Young (1995) enables the measurement of living-fungal mass in samples of naturally decaying saltmarsh-grass blades beneath as little as 5 mm" leaf-surface area (1 ng of ergosterol with 50 ~11 injections in HPLC, equivalent to 0.2 1-G of organic fungal mass) (Fell and Newell, 1998). Ergosterol is readily and fully separable from other common sterols (brassicasterol, campesterol, cholesterol, fucosterol, sitosterol, stigmasterol) using liquid chromatography (Newell and Fallon, 199l). Newell and Fallon (1991; Newell, 1993a) expanded the power of the ergosterol method by providing it with the capability for measurement of rates of production of fungal mass. This expansion involved the incubation of decaying-plant samples with radiolabeled acetate (the basic, smallmolecular precursor in ergosterol synthesis: Mercer, 1984; Gessner and Newell, 2001), subsequently separating and measuring the amount of ergosterol in the samples by liquid chromatography, capturing the ergosterol peak during chromatography, measuring the extent to which the ergosterol peak was made radioactive (= the degree of incorporation of acetate) during the incubations, and calculating the rate of production of organic fungal mass (Gessner and Newell, 2001). Acetate was chosen as the tagged precursor because of the possibility that precursors closer to 359
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the final product (e.g. mevalonate) would not be readily taken u p from outside of the cell (see Serrano-Carreon et al., 1993). See Arao (1999) for description of an alternative m e t h o d for monitoring acetate incorporation into fungal lipids.
Detailed methodology Equipment and reagents I include here and in the next subsection the core materials and instructions needed for acetate incubations and ergosterol analyses. See Newell (1993a; 1996a; 2000) for additional details, including methods for obtaining parallel organic densities of natural samples. USE CAUTION: consult local radiation-safety officials for rules for handling radioisotopes and radioactive waste; always work in fume hoods tested for effective function and wear protective clothing w h e n handling radioisotopes and solvents; remember that solvents are highly flammable (especially pentane) - - remove all flame and spark sources. • Carbon-14 acetate (I-['4C]acetate), sodium salt, crystalline solid, specific activity 1-10mCimmol' (37-370 NBq mmol ') (e.g. ICN #12014); store tightly sealed at 0-5°C - - it is hygroscopic and efflorescent and will sublime. The crystalline solid form is recommended here, because the dissolved form is provided in ethanol, which one must remove if one desires to avoid adding ethanol to samples. • Hultiplace filtration manifold for rinsing ['4C]-incubated samples (these can be constructed in-house for much less than scientific-supply prices). • Pure ergosterol for standards; store desiccated under ]'q2 at 0-5°C, or oxidation (can be seen as yellowing) will occur (Newell, 1994b). • HPLC-grade solvents in repipetting containers: methanol, reagent ethanol, pentane; scintillation fluor. • Potassium hydroxide and anhydrous sodium acetate. • Refluxing system (multiple reflux units in an 80°C water bath, plus a source of cooling water with tubing connections to the condensers); grease-free, screw-thread, Teflon connectors (Wheaton Connections®) make this system considerably more user-friendly than condensers with ground-glass connectors. • Dry bath, glass vials to fit bach compartments snugly, and drying-gas distribution manifold. • Sonicating cleaning bath for redissolutions of dried-down neutral lipids. • High-performance liquid chromatographic (HPLC) system (pump, injection valve, column, UV detector [at least 282 and 210 nm detection; 282 for ergosterol, 210 for other sterols], integrator/recorder), with 2-way valve at detector exit (all parts compatible with methanol). • Gas/liquid-tight syringe (5001ul; Teflon plunger) to match injection valve receiver fitting (e.g. luer lock). • Scintillation counter.
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Incubation and assay
1. Place standard samples (e.g. pieces of marshgrass leaf blades of measured surface area, cut from blades, rinsed and stored air-dry [Newell et al., 2000]) in sterile, clean 20 ml, screw-cap vials. Submerge each sample in a measured quantity of bacteria-free seawater (0.2-1amor, ideally, 0.1-1am-filtered; Kirchman and Ducklow, 1993; Blosse et al., 1998; Gasol and Moran, 1999; do not allow filters to run dry: Kiene and Linn, 1999). Cap loosely. Bring two (or more) replicate sample vials to 2% formaldehyde and cap tightly (killed controls). 2. Incubate vials under controlled conditions of temperature and light with slow agitation (e.g. -< 60-75 reciprocations per min through 0.5 cm; violent agitation can have negative effects on fungi: Newell et al., 1996) for 1-3 h to permit adaptation of fungi to submerged conditions (Newell et al., 1985). Slow agitation can be achieved by placing vials loosely in a rack on a rotary shaker run at its slowest maintainable speed, or purchase a slow-speed shaker (see List of suppliers). 3. Add [1-'4C]acetate plus non-radioactive sodium acetate in bacteriafree aqueous solution to bring each incubation vial to 5 mM acetate. Pilot experimentation is needed to find appropriate ratios of radioactive and non-radioactive acetate (Newell, 1993a). Add measured quantity of non-radioactive acetate powder to original vial of radioactive acetate, and add measured volume of bacteria-free water, calculated to be the appropriate amount for micropipetting as aliquots to incubation vials. 5 mM acetate has been repeatedly found to maximize incorporation of acetate into ergosterol (Gessner and Newell, 2001), but ideally, pilot experiments should be run to test whether a particular type of sample conforms to this generalization. Record time of initiation of incubations. Incubate as above for the pre-adaptations for 0.25-4 h (depending on rate of incorporation of acetate, until CPM in ergosterol is clearly measurable, but not long enough to move out of the linearrate period; pilot experiments needed here) (Newell and Fallon, 1991; Gessner and Newell, 2001 ). 4. Terminate acetate incubations by rinsing x2 in bacteria-free seawater (volume = x4 incubation volume) (Suberkropp and Weyers, 1996; fixation and storage at this point can cause loss of label: Newell and Fallon, 1991). Record time of terminations. Immediately place rinsed samples in 5 ml reagent ethanol and store at 4°C in darkness. 5. Extract ergosterol by methanol refluxing and pentane partitioning as described in detail in Newell (2000; Newell, 1993a). This includes: refluxing for 2 h in methanol; addition of KOH/ethanol and 30 min refluxing (lysis of steryl esters); addition of H~O and partioning into pentane (separation from fatty acids and other polar molecules); drying down of pentane in dry bath; redissolution in methanol (cleaning-bath sonication); passage through 0.45 1ira filters into 2 ml, Teflon-capped glass vials. Include procedural standards in the extraction process (add pure ergosterol solutions in methanol to reflux flasks of methanol at a range of final concentrations encompassing those expected from samples, and take them through the whole procedure). 361
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Ideally, one should also add spikes of ergosterol to split natural samples (Newell, 1993a; 2000) to check for sample effects upon ergosterol recovery. See Young (1995) or Eash et al. (1996) for alternative extraction procedures involving microwaving or ultrasonication rather than refluxing. Run samples through HPLC with methanol eluant (Newell, 2000). Inject sufficient sample into injection valve (3 to 4 times injection-loop volume) to ensure that injection loop is sample-rinsed and full of sample (beware of dead volume ahead of injection loop; Newell, 2000; Gessner and Newell, 2001). Inject freshly prepared ergosterol solutions at a concentration in the mid-range of sample concentrations as injection standards (comparison to procedural standards permits calculation of losses of ergosterol during the assay). At 2 ml min ~, 282 nm detection, reverse-phase column (220 x 4.6 mm, 10 1Am), all peaks appear within about 10 min (Newell, 2000). Rinse syringe and injection valve with methanol between samples. During the chromatography of injection standards, record the time of the beginning and ending of the ergosterol peak as shown on the integrator/recorder. During chromatography of samples, at the moment the sample-ergosterol peak appears, open the 2-way valve at the detector exit so that eluant flow is to the open vent (rather than to the vent leading to the waste bottle). Hold a clean scintillation vial under the open vent and collect the sample ergosterol peak. Turn the 2-way valve back to waste just prior to the end of the ergosterol peak. Rinse 2-way valve between samples by allowing clean methanol eluant to flow through the open vent. Collect standard peaks to use to obtain background CPM (scintillation counts per minute) values. Add 10 ml scintillation fluor to vials containing ergosterol peaks. Agitate to mix fluor with methanol, and count vials in a scintillation counter. Subtract counts exhibited by killed controls from live-sample counts to find counts due to fungal incorporation. Find ergosterol concentrations per sample by comparison of sample peak areas with peak areas for procedural standards. Use the CPM for samples, minus tlle background CPM, and the equation of Newell (1993a) to find specific rate (~L, day ~) of ergosterol synthesis, but multiply acetate specific activity (SA) by 0.89 to adjust for losses of carbon atoms when using [l-"C]acetate (Newell, 2000; Gessner and Newell, 2001). Alternatively, use an empirically determined conversion factor to calculate rates of fungal production (Gessner and Newell, 2001; note especially that the conversion factors of Newell (1996b) are high due to a consistent peculiarity of the HPLC injection valve used - - adjust these factors downward by x0.65; the resultant value for the ascomycetes of smooth cordgrass is 12.6 ~g fungal organic mass per nmol acetate incorporated into ergosterol).
Troubleshooting An overriding issue to keep in mind in designing research projects with the goal of studying decay of vascular plants, is that one must avoid
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subjecting the substrate to unnatural conditions that would very likely result in artifactitious experimental outcomes. Severing from the shoot of non-abscised parts, oven-drying, grinding, using green, living shoots or leaves as initial material, litterbagging on sediment surfaces, and permanent submergence of substrates are potentially contraindicated procedures; see Newell (1993b, 1996a), B~rlocher (1997), Gessner et al. (1999). Since ergosterol is a part of the plasma membrane of living hyphal cells, it is susceptible to lysis at cell death. Therefore, sample-preservation methods that slowly kill the fungi within decaying substrates should be avoided. One of these methods is oven-drying (Newell, 2000). For standing-decaying saltmarsh-grass shoots, alternating periods of wetness and dryness are natural, and the fungi of this decay system are not set back by air-drying, so this is an effective method of sample preservation (Newell el al., 2000). This may not be true, however, for samples of persistently submerged leaves such as those of mangroves or riparian leaves in freshwater streams (Gessner and Newell, 2001). gjurman (1994) found that a fungal species that had been grown on wood under damp chamber conditions lost 93% of its ergosterol when humidity was reduced and the wood became desiccated. The safest approach is to move samples directly to methanol or ethanol submergence (Newell, 2000). Losses of ergosterol during extraction and chromatography are characteristically small (< 15~/,; Gessner and Newell, 2001), but there exists the possibility that individual batches could suffer larger losses (accidental temperature shifts, unanticipated interactions among sample chemicals, etc.). Therefore, it is essential that procedural standards be run (see assay step 5), rather than simply standardizing chromatographic runs with a set of injection standards that did not experience the whole extraction~chromatography impact. Although a general conversion factor for calculating fungal organic mass from ergosterol content of decaying substrates has been proposed (200 units organic mass per unit ergosterol; Newell, 2000; Gessner and Newell, 2001), the range of potential conversion factors is large, based on reported ergosterol contents of the fungal species tested (Gessner and Newell, 2001). It is possible that much of this variation is due to genetic change following adaptation to 'luxury' culture conditions during isolate maintenance and transfer. Note that rich culture media can induce low ergosterol contents (Newell, 1992; Bjurman, 1994; Gunnarsson et al., 1996), and consider, for example, the clear cases of constitutive culturinginduced change from the natural state found by Gramss (1991) for wooddecay fungi. When five species of saltmarsh-grass ascomycetes from two distinct major taxa and two decomposition systems were tested immediately upon capture from nature, the five had homogeneous mycelialergosterol content (CV = 8%) only 25~5 different from the proposed general average of 5 mg ergosterol g ' organic fungal mass (Newell, 2000). The empirical conversion factors for calculation of rates of fungal production from rates of incorporation of acetate into ergosterol currently range widely (about 6-19 lJg fungal organic mass per nmol acetate incorporated) (Gessner and Newell, 2001). These values are about 20% to greater than 6-fold the theoretical conversion factor, implying that it is not 363
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possible to force the fungi, in the empirical situations that have been tried, to use offered, radiolabeled precursor exclusively, a situation analogous to that found for bacterial-productivity methods (Jeffrey and Paul, 1988). Only two conversion factors are currently available for marine substrates, 12.6 lJg fungal organic mass per nmol acetate for smooth cordgrass (Spartina alterniflora) and 17.8 lag nmol ~ for black needlerush (]uncus roemerianus) (adjusted from Newell, 1996b; Gessner and Newell, 2001). The needlerush value is very tenuous, since it is based on data for only one species (versus four species for the cordgrass value). Much more work is needed to establish the true range of this conversion factor, and the causes of its variability. See Gessner and Newell (2001) for further discussion of potential problems involved in the acetate-to-ergosterol method.
Application Estimation of fungal-mass content using ergosterol as a proxy has become a standard technique, not only in ecological work but also in agricultural science, plant pathology, industrial applications, and food science (Gessner, 1997; Miller and Young, 1997; Nout et al., 1997; Pereira et al., 1998; Dowell et al., 1999; Joergensen and Scheu, 1999; Kuehn et aI., 1999; Schn6rer et al., 1999; Marstorp et al., 2000; Sridhar and B/irlocher, 2000). In marine science, the ergosterol biomass method has been used extensively in the saltmarsh ecosystem (review: Newell and Porter, 2000; see also Castro and Freitas, 2000), and sparingly in the mangrove ecosystem and on wood decaying in a marine system (Miller et al., 1985; Newell and Fell, 1992). One surprising result of this work is that the saltmarsh and mangrove ecosystems appear to be very different with regard to the major microbial types driving leaf decay: in saltmarshes, ascomycetous fungi accumulate substantial biomass (10-20% or more of decaying-system organic mass) as they decompose standing-decaying leaf blades, but in mangroves, fungi exhibit negligibly small biomasses in submerged decaying mangrove leaves - - oomycotes and bacteria probably do most of this submerged-leaf digestion (Newell, 1996a). Application of the acetate-to-ergosterol method has been much more limited than use of ergosterol as a biomass proxy. Principal ecosystems in which the acetate-to-ergosterol method has been used have been saltmarshes, freshwater marshes, and freshwater streams (Newell et al., 1995; Gessner, 1997; Suberkropp, 1998; Newell and Porter, 2000; Kuehn et al., 2000). For the marine, saltmarsh system, several unexpected results have been obtained. For example: (i) fungal (ascomycetous) productivity per m: of marsh per day in Georgia (USA) marshes has been preliminarily estimated to be about one-half as great as that of total bacteria (bacteria mostly in sediment, to 20 cm depth) per m a marsh in summer, but about xl0 greater than total bacteria in winter (Newell and Porter, 2000; cf. Castro and Freitas, 2000); (ii) rates of fungal production per unit decaying marshgrass blade can be as high (at a standard temperature of 20°C) in northerly marshes (Wells Reserve, Maine, 43°N) as they are in southtemperate marshes (three Florida marshes, 29-30°N) (Newell et al., 2000).
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Many other surprises are likely as the acetate-to-ergosterol method finally enables measurement of fungal productivity in further marine-oriented and other laboratories (Pennanen et al., 1998; Reid et al., 1999). "o ¢.
CONCLUSION The ergosterol biomass-proxy method is a robust one, having been repeatedly challenged and cross-checked with other methods, and having held up well (Newell, 2000; Gessner and Newell, 2001). The acetate-to-ergosterol method of measuring fungal productivity could use more attention from a broader range of scientists to improve circumscription of conversion factors and understanding of other factors affecting the method's accuracy (Newell and Porter, 2000; Gessner and Newell, 2001). There is still a great deal of room for application of the ergosterol-based biomass and productivity methods at the oceans' edges, and in freshwater and terrestrial ecosystems, in order to develop a thorough grasp of patterns of fungal standing crops and rates of fungal throughput.
FUTURE DIRECTIONS Both living and total (empty-hyphal plus cytoplasm-filled hyphal mass) fungal mass could and probably should be measured in parallel (Newell, 1996a) whenever practicable (Ekblad et al., 1998). It may be that these two fungal indices could be measured in one chromatographic run, perhaps by derivatizing either or both glucosamine and ergosterol (Ikemoto et al., 1992), so that they could be measured during one reverse-phase HPLC assay with UV detection (Osswald et al., 1995) (though extractions would still have to be separate and parallel). This combination assay could be made more powerful by the addition of acetate-to-ergosterol application, along with an analogous glucosamine-to-chitin ramification (Roff et al., 1994) or perhaps even an increasingly-fluorescent-chitin method of measuring fungal productivity (Carrano et al., 1997). The cost of [1-1~C]acetate is a hindrance to analyses of large numbers of replicate samples in the acetate-to-ergosterol method (the 1-C-labeled version [ICN/2000 catalog, about $360 mCi '] is 0.4-0.5-fold less costly than the 2-C and 1,2-C versions). It is possible that tritiated acetate could be routinely substituted for [1-'~Clacetate, and the cost ratio, on a mCi basis (1 mCi = 37 MBq), between the two radiochemicals is about 8:1, HC:~H, at the time of writing (ICN catalog for 2000). Newell and Fallon (1991) compared tritiated acetate and [1-"Clacetate in the original acetateto-ergosterol work, and found that pure cultures of two saltmarsh ascomycetes would incorporate measurable quantities of tritium into ergosterol, but the use of tritiated acetate in the acetate-to-ergosterol method has not been further tested. The halophytophthoras are marine-restricted, mycelial eukaryotic organoosmotrophs in the phylum Oomycota, Kingdom Straminipila (see Introduction). They are involved in the decay of leaves that fall into the 365
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marine environment (Newell and Fell, 1997, and references therein), and their zoospores are a part of the coastal-marine plankton (Newell and Fell, 1996). There are no published methods for measurement of biomass or productivity of oomycotes (they do not synthesize ergosterol, and most do not synthesize chitin) (Fell and Newell, 1998). One potential means of measuring h a l o p h y t o p h t h o r a n standing crops might be to use commercially available immunodiagnostic kits designed for testing for the presence of plant-pathogenic species of Phytophthora, which are closely related to the Halophytophthora spp. (Ali-Shtayeh et aI., 1991; Miller et al., 1997; Werres et al., 1997). There m a y well be enough breadth in immunospecificity (Gabler and Richter, 1997; Miller et al., 1997; Wakeham et al., 1997) that the commercial kits could be used in immunoassays for halophytophthoran biomass. Productivity of halophytophthoras might also be measurable analogously to the acetate-to-ergosterol m e t h o d for fungi - the radioprecursors would be to cell-wall glucans not synthesized by fungi (Fell and Newell, 1998) or to the oomycotic phospholipid fatty acid 20:5 (Gessner and Newell, 2001). Gessner and Newell (2001) provide other examples of probable future directions in the evolution of the ergosterol/fungal methods, including the marriage of these non-species-specific methods to DNA-technological or i m m u n o a s s a y methods that are highly species-specific (such as competitive PCR, FISH, and ELISA; Clausen, 1997; Dewey et al., 1997; Nicholson et al., 1998; Kessel et al., 1999; Spear et al., 1999).
References Alexopoulos, C. J., Mims, C. W. and Blackwell, M. (1996). hztroductory Mycology, 4th edn. Wiley, New York. Ali-Shtayeh, M. S., MacDonald, J. D. and Kabashima, J. (1991). A method for using commercial ELISA tests to detect zoospores of Phytophthora and Pythiunl in irrigation water. Plant Disease 75, 305-311. Arao, T. (1999). In situ detection of changes in soil bacterial and fungaI activities by measuring 13C incorporation into soil phospholipid fatty acids from 13C acetate. Soil Biol. Biochem. 31, 1015-1020. B~irlocher, E (1997). Pitfalls of traditional techniques when studying decomposition of vascular plant remains ill aquatic habitats. Limnetica 13, 1-11. Beakes, G. W. (1998). Relationships between lower fungi and protozoa. In: Evolutionary Relationships Amollg Protozoa (G, H. Coombs, K. Vickerman, M. A. Sleigh and A. Warren, Eds), pp. 351-373. Chapman & Hall, London. Bjurman, J. (1994). Ergosterol as an indicator of mould growth on wood in relation to culture age, humidity stress and nutrient level, hzt. Biodeterior. Biod~,grad. 33, 355-368. Blosse, P. T., Boulter, E. M. and Sundaram, S. (1998). Diminutive bacteria: implications for sterile filtration. Amer. Biotechnol. Lab. 16, 38-40. Carrano, L., Tavecchia, P., Sponga, E and Spreafico, E (1997). Dansyl N-acetyl glucosamine as a precursor of fluorescent chitin - - a method to detect fungal cell-wall inhibitors. J. Al#ibiotics 50, 177-179. Castro, P. and Freitas, H. (2000). Fungal biomass, senescence and decomposition in Spartilza maritima in tile Mondego salt marsh (Portugal). Hydrobiologia, 428, 171-177.
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Clausen, C. A. (1997). Immunological detection of wood decay fungi --- an overview of techniques developed from 1986 to the present. Int. Biodeterior. B iodegrad. 39, 133-143. DeLong, E. E (1998). Molecular phylogenetics: new perspective on the ecology, evolution and biodiversity of marine organisms. In: Molecular Approaches to the Study of tire Oceans (K. E. Cooksey, Ed.), pp. 1-27. Chapman & Hall, London. Dewey, F. M., Thornton, C. R. and Gilligan, C. A. (1997). Use of monoclonal antibodies to detect, quantify and visualize fungi in soils. Adv. Bot. Res. 24, 275-308. Dick, M. W., Vick, M. C., Gibbings, J. G., Hedderson, T. A. and Lopez Lastra, C. C. (1999). 18S rDNA for species of Leptoh\\~nia and other Peronosporomycetes: justification for the subclass taxa Saprolegniomycetidae and Peronosporomycetidae and division of the Saprolegniaceae seusu lato into the Leptolegniaceae and Saprolegniaceae. Mycol. Res. 103, 1119-1125. Djajakirana, G., Joergensen, R. G. and Meyer, B. (1996). Ergosterol and microbial biomass relationship in soil. Biol. Fertil. Soils 22, 299-304. Dowell, E E., Ram, M. S. and Seitz, L. M. (1999). Predicting scab, vomitoxin, and ergosterol in single wheat kernels using near-infrared spectroscopy. Cereal Chem. 76, 573-576. Eash, N. S., Stahl, P. D., Parkin, T. B. and Karlen, D. L. (1996). A simplified method for extraction of ergosterol from soil. Soil Sci. Soc. Am. J. 60, 468-471. Ekblad, A., Wallander, H. and N~isholm, T. (1998). Chitin and ergosterol combined to measure total and living fungal biomass in ectomycorrhizas. New Phytol. 138, 143-149.
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compounds containing hydroxyl and amino groups with the fluorogenic reagent, 2-naphthoylimidazole. Angew. Chem., Int. Ed. Engl. 31, 890-891. Jeffrey, W. H. and Paul, J. H. (1988). Underestimation of DNA synthesis by l~H]thymidine incorporation in marine bacteria. Appl. Environ. Microbiol. 54, 3165-3168. Joergensen, R. G. and Scheu, S. (1999). Depth gradients of microbial and chemical properties in moder soils under beech and spruce. Pedobiologia 43, 134-144. Kendrick, B. (1992). The Fifth Kingdom, 2nd edn. Focus Information Group, Newburyport, MA. Kessel, G. J. T., de Haas, B. H., Lombaers-van der Plas, C. H., Meijer, E. M. J., Dewey, E M., Goudriaan, J., van der Werf, W. and K6hl, J. (1999). Quantification of mycelium of Botrytis spp. and the antagonist LUocladium atrum in necrotic leaf tissue of cyclamen and lily by fluorescence microscopy and image analysis. Phytopatholo~cy 89, 868-876. Kiene, R. P. and Linn, L. J. (1999). Filter type and sample handling affect determination of organic substrate uptake by bacterioplankton. Aquatic Microbial Ecol. 17, 311-321. Kirchman, D. L. and Ducklow, H. W. (1993). Estimating conversion factors for the thymidine and leucine methods for measuring bacterial production. In: Handbook of Methods in Aquatic Microbial Ecology (P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 579-586. Lewis Publishers, Boca Raton, FL. Kohlmeyer, J. and Volkmann-Kohlmeyer, B. (1991). Illustrated key to the filamentous higher marine fungi. Bot. Mar. 34, 1-6l. Kohlmeyer, J., Volkmann-Kohlmeyer, B. and Eriksson, O. E. (1999). Fungi on Juncus tvemerianus 12. Two new species of Mycosphaerella and Paraphaeosphaeria (Ascomycotina). Bot. Mar. 42, 505-511. Kuehn, K. A., Gessner, M. O., Wetzel, R. G. and Suberkropp, K. (1999). Decomposition and CO~ evolution from standing litter of the emergent macrophyte Erh~nthus gi~anteus. Microbial Ecol. 38, 50-57. Kuehn, K. A., Lemke, M. J., Suberkropp, K. and Wetzel, R. G. (2000). Microbial biomass and production associated with decaying leaf litter of the emergent macrophyte ]uncus effusus. Limnol. Oceanogr. 45, 862-870. Lee, C., Howarth, R. W. and Howes, B. L. (1980). Sterols in decomposing Spartina altern(flora and the use of ergosterol in estimating the contribution of fungi to detrital nitrogen. Limnol. Oceanogr. 25, 290-303. Marstorp, H., Guan, X. and Gong, P. (2000). Relationship between dsDNA, chloroform labile C and ergosterol in soils of different organic matter contents and pH. Soil Biol. Biochem. 32, 879-882. Mercer, E. I. (1984). The biosynthesis of ergosterol. Pesticide Sci. 15, 133-155. Miller, J. D. and Young, I. C. (1997). The use of ergosterol to measure exposure to fungal propagules in indoor air. Amer. lndustr. Hyg. J. 58, 39-43. Miller, J. D., Jones, E. B. G., Moharir, Y. E. and Finlay, J. A. (1985). Colonization of wood blocks by marine fungi in Langstone Harbour. Bot. Mar. 28, 251-257. Miller, S. A., Madden, L. V. and Schmitthenner, A. E (1997). Distribution of Phytophthora spp. in field soils determined by immunoassay. Phytopathology 87, 101-107. Newell, S. Y. (1992). Estimating fungal biomass and productivity in decomposing litter. In: The Fungal Community, 2nd edn (G. C. Carroll and D. T. Wicklow, Eds), pp. 521-561. Marcel-Dekker, New York. Newell, S. Y. (1993a). Membrane-containing fungal mass and fungal specific growth rate in natural samples. In: Handbook qf Methods in Aquatic Microbial Ecolo??y(P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 579-586. Lewis Publishers, Boca Raton, FL.
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Newell, S. Y. (1993b). Decomposition of shoots of a saltmarsh grass; methodology and dynamics of microbial assemblages. Adv. Microbial Ecol. 13, 301-326. Newell, S. Y. (1994a). Ecomethodology for organoosmotrophs: prokaryotic unicellular versus eukaryotic mycelial. Microbial Ecol. 28, 151-157. Newell, S. Y. (1994b). Total and free ergosterol in mycelia of saltmarsh ascomycetes with access to whole leaves or aqueous extracts of leaves. Appl. Environ. Microbiol. 60, 3479-3482. Newell, S. Y. (1996a). Established and potential impacts of eukaryotic mycelial decomposers in marine/terrestrial ecotones. J. Exp. Mar. Biol. Ecol. 200, 187-206. Newell, S. Y. (1996b). The [HC]acetate-to-ergosterol method: factors for conversion from acetate incorporated to organic fungal mass synthesized. Soil Biol. Biochem. 28, 681-683. Newell, S. Y. (2000). Methods for determining biomass and productivity of mycelial marine fungi. In: Marine Mycology - - A Practical Approach (K. D. Hyde and S. B. Pointing, Eds), pp. 69-91. Fungal Diversity Press, Hong Kong. Newell, S. Y. and Fallon, R. D. (1991). Toward a method for measuring instantaneous fungal growth rates in field samples. Ecology 72, 1547-1559. Newell, S. Y. and Fell, J. W. (1992). Ergosterol content of living and submerged, decaying leaves and twigs of red mangrove. Can. J. Microbiol. 38, 979-982. Newell, S. Y. and Fell, J. W. (1996). Cues for zoospore release by marine oomycotes in naturally decaying submerged leaves. Mycologia 88, 934-938. Newell, S. Y. and Fell, J. W. (1997). Competition among mangrove oomycotes, and between oomycotes and other microbes. Aquatic Microbial Ecol. 12, 21-28. Newell, S. Y. and Porter, D. (2000). Microbial secondary production from saltmarsh-grass shoots, and its known and potential fates. In: Concepts and Controversies in Tidal Marsh Ecology (M. P. Weinstein and D. A. Kreeger, Eds), 159-185. Kluwer, Amsterdam. Newell, S. Y. and Wall, V. D. (1998). Response of saltmarsh fungi to the presence of mercury and polychlorinated biphenyls at a Superfund site. Mycolo~ia 90, 777-784. Newell, S. Y., Fallon, R. D., Cal Rodriguez, R. M. and Groene, L. C. (1985). Influence of rain, tidal wetting and relative humidity on release of carbon dioxide by standing-dead salt-marsh plants. Oecologia 68, 73-79. Newell, S. Y., Moran, M. A., Wicks, R. and Hodson, R. E. (1995). Productivities of microbial decomposers during early stages of decomposition of leaves of a freshwater sedge. Freshwater Biol. 34, 135-148. Newell, S. Y., Porter, D. and Lingle, W. L. (1996). Lignocellulolysis by ascomycetes (Fungi) of a saltmarsh grass (smooth cordgrass). Microsc. Res. Technol. 33, 32--46. Newell, S. Y., Blum, L. K., Crawford, R. E., Dai, T. and Dionne, M. (2000). Autumnal biomass and potential productivity of saltmarsh fungi from 29 ° to 43 ~' north latitude along the United States Atlantic coast. Appl. E~lviro17. Microbiol. 66, 180-185. Nicholson, P., Simpson, D. R., Weston, G., Rezanoor, H. N., Lees, A. K., Parry,, D. W. and Joyce, D. (1998). Detection and quantification of Fusarium culmorum and Fusarium gramillcnrllm in cereals using PCR assays. Physiol. Mol. PlaHt Pathol. 53, 17-37.
Nout, M. J. R., Rinzema, A. and Smits, J. P. (1997). Biomass and productivity estimates in solid substrate fermentations. In: The Mycota IV. E~lviromuental altd Microbial Rehltio~iships (D. T. Wicklow and B. S6derstr6m, Eds), pp. 323-345. Springer Verlag, New York. Osswald, W. F., Jehle, J. and Firl, J. (1995). Quantification of fungal infection in plant tissues by determining the glucosamine phenylisothiocyanate derivative using HPLC techniques. J. Plaut Physiol. 145, 393-397.
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Pennanen, T., Frieze, H., Vanhala, E, Kiikkil5, O., Neuvonen, S. and Bfi•th, E. (1998). Structure of a microbial community in soil after prolonged addition of low levels of simulated acid rain. Appl. Environ. Microbiol. 64, 2173-2180. Pereira, A. P., Graqa, M. A. S. and Molles, M. (1998). Leaf litter decomposition in relation to litter physico-chemical properties, fungal biomass, arthropod colonization, and geographical origin of plant species. Pedobiolosia42, 316-327. Reid, L. M., Nicol, R. W., Ouellet, T., Savard, M., Miller, J. D., Young, J. C., Stewart, D. W. and Schaafsma, A. W. (1999). Interaction of Fusarium graminearum and E moniliforme in maize ears: disease progress, fungal biomass, and mycotoxin accumulation. Phytopathology 89, 1028-1037. Roff, J. C., Kroetsch, J. T. and Clarke, A. J. (1994). A radiochemical method for secondary production in planktonic crustacea based on rate of chitin synthesis. J. Plankton Res. 16, 961-976. Schn~rer, J., Olsson, J. and B6rjesson, T. (1999). Fungal volatiles as indicators of food and feeds spoilage. Fungal Genet. Biol. 27, 209-217. Seitz, L. M., Sauer, D. B. Burroughs, R., Mohr, H. E. and Hubbard, J. D. (1979). Ergosterol as a measure of fungal growth. Phytopathology 69, 1202-1203. Serrano-Carreon, L., Hathout, Y., Bensoussan, M. and Belin, J.-M. (1993). Metabolism of linoleic acid or mevalonate and 6-pentyl-R-pyrone biosynthesis by Trichoderma species. Appl. Environ. Mic1~biol. 59, 2945-2950. Spear, R. N., Li, S., Nordheim, E. V. and Andrews, J. H. (1999). Quantitative imaging and statistical analysis of fluorescence in situ hybridization (FISH) of Aureobasidium pullulans. J. Microbiol. Methods 35, 101-110. Sridhar, K. R. and BSrlocher. (2000). Initial colonization, nutrient supply, and fungal activity on leaves decaying in streams. Appl. Environ. Microbiol. 66, 1114-1119. Suberkropp, K. (1998). Microorganisms and organic matter decomposition. In: River Ecology and Management (R. J. Naiman and R. E. Bilby, Eds), pp. 120-143. Springer, New York. Suberkropp, K. and Weyers, H. (1996). Application of fungal and bacterial production methodologies to decomposing leaves in streams. Appl. Environ. Miclvbiol. 62, 1610-1615. Wakeham, A. J., Pettitt, T. R. and White, J. G. (1997). A novel method for detection of viable zoospores of Pythium in irrigation water. Ann. Appl. Biol. 131, 427-435. Weete, J. D., Fuller, M. S., Huang, M. Q. and Ghandi, S. (1989) Fatty acids and sterols of selected hyphochytridiomycetes and chytridiomycetes. Exp. Mycol. 13, 183-195. Werres, S., Hahn, R. and Themann, K. (1997). Application of different techniques to detect Phytophthora spp. in roots of commercially produced Chamaecyparis lawsoniana. J. Plant Disease Protect. 104, 474-482. Young, J. C. (1995). Microwave-assisted extraction of the fungal metabolite ergosterol and total fatty acids. J. A~ric. Food Chem. 43, 2904-2910.
List of suppliers The firms b e l o w are listed b e c a u s e of the a u t h o r ' s familiarity w i t h them, not b e c a u s e they are necessarily the best sources of the m a t e r i a l s that they sell. There is c o n s i d e r a b l e p r o d u c t o v e r l a p a m o n g several of the c o m p a nies b e l o w - - m a n y sell several of the items listed here.
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Alltech Associates 2051 Waukegan Road Deerfield, IL 60015, USA www.alltechweb.com Tel: 847-948-8600 Fax: 847-948-1078
Hamilton Company 4970 Energy Way Reno, N V 89520, USA www.hamiltoncompany.com Tel: 800-648-5950 Fax: 775-856-7259
HPLC gear, including 2-way valves
Gas/liquid-tight syringes ICN Biomedicals 3300 Hyland Avenue Costa Mesa, CA 92626, USA ~ww.icnbionled.co~n Tel: 800-854-0530 Fax: 800-334-6999
Anspec/Ohio 12 W Selby Blvd., Suite 4 Columbus, OH 43085 www.munhall-anspec.com Tel: 800-521-1720 Fax: 614-888-7843
[~4C]acetate
Valves, tubing for HPLC Kimble/Kontes 1022 Spruce Street Vineland, N] 08360, USA www.kimble-kontes.com Tel: 888-546-2351 Fax: 609-692-3242
Beckman Coulter PO Box 3100 Fullerton, CA 92834, USA www.becklIla71co~lter.com Tel: 714-773-6707 Fax: 714-773-8186
HPLC fittings, tubing
Scintillation counters LC.GC Magazine PO Box 6168 Duluth, M N 55806, USA www.lcgcmag.com Teh 888-527-7008
Dionex Corporation 1228 Titan Way Sunnyvale, CA 94086, USA www.dionex.com Tel: 800-723-1161 Fax: 408-739-4398
Contact info. for chromatography suppliers (subscription gratis)
HPLC apparatus N e w Brunswick Scientific PO Box 4005 Edison, NJ 08818, USA
Fisher Scientific 585 Alpha Drive Pittsburgh, PA 15238, USA
WWW.IIbSC.C~}tl
www.fishersci.cotH
Teh 800-631-5417 Fax: 908-287-4222
Teh 800-766-7000 Fax: 800-926-1166
Slow-speed shakers (innova)
Inexpensive HPLC apparatus
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PE Biosystems
Supelco
761 Main Avenue Norwalk, CT 06859, USA www.perkin-elmer.com Teh 203-762-4000 Fax: 203-762-4228
Supelco Park Bellefonte, PA 16823, USA www.sigma-aldrich.com Tel: 800-247-3010 Fax: 800-325-5052
UV detector for HPLC
HPLC columns; other HPLC items
Phenomenex
Wheaton Scientific Products
2320 W. 205th Street Torrance, CA 90501, USA www.phenomenex.com Teh 310-212-0555 Fax: 310-328-7768
1501 N lOth Street Millville, NJ 08332, USA www.wheatonsci.com Teh 800-225-1437 Fax: 609-825-1368
HPLC columns
Grease-free reflux glassware
Sigma-Aldrich PO Box 14508 St. Louis, MO 63178, USA www.sigma-aldrich.com Teh 800-325-3010 Fax: 800-325-5052
Ergosterol, other chemicals
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19 Molecular Phylogeny: Applications and Implications for Marine Microbiology c-
Craig L Moyer Department of Biology, MS#9160, Western Washington University,Bellingham,Washington 98225, USA
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CONTENTS Introduction to the application of molecular phylogenyto marine microbiology Methodology for the generation and analysis of SSU rDNA clone libraries Methodology for the generation and phylogeneticanalysisof SSU rDNA sequences Concluding remarks
I N T R O D U C T I O N T O T H E A P P L I C A T I O N OF MOLECULAR PHYLOGENYTO MARINE MICROBIOLOGY The field of marine microbiology has from its inception been a methodslimited proposition, whether microbial communities are characterized through an autecology or synecology perspective. When the focus has been towards autecology, or the characterization of microbial populations through the study of cultured isolates and their physiology, the approach encompasses microbial growth procedures, such as dilution to extinction methods or enrichment culture. The primary limitation continues to be the frequent dependence upon nutrient-laden media to satisfy the nutritional requirements of every population of microorganisms which exists within the community. 'The most one can hope for is a medium in which many microorganisms will grow and with which the results may be duplicated' (ZoBell, 1946). The overall goal is to understand how microbial populations are able to adapt to a range of environmental parameters (or limitations) and yet influence marine microbiological processes. For a review of autecological studies emphasizing the predominant forcing functions (e.g. salinity, temperature, hydrostatic pressure, and nutrient availability) of the marine environment and their impact on microorganisms, see Morita (1986). Synecology or a systems-level 'black box' approach towards studying an entire community employs the central tenant that emergent properties result from the organization of the whole METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0 12-521530 4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any form reserved
community which would otherwise be unobserved (i.e. the whole is greater than the sum of the parts). This general approach uses methods that estimate the in silu microbial biomass, viability, metabolism and growth through deterministic assays of environmental samples. For example, the most common strategy used to enumerate the total number of microorganisms present (i.e. biomass) in a marine sample relies on direct microscopic counts, which lacks any capability for differentiation beyond simple morphology. For a detailed review of the marine microbiological methodology used in predominantly synecological studies, see Karl (1986). A suite of molecular biological methods revolving around the idea that cellular component analyses provide a culture-independent means of investigating microorganisms as they occur in nature was developed in the mid-1980s (Olsen et al., 1986; Pace et al., 1986). This methodological approach targets a microbial community's primary members through molecular (i.e. cell component) means and characterizes their respective phylogeny or evolutionary history. Over the last decade, numerous studies using these molecular biological approaches have significantly changed our understanding of marine microbiology, fueling new avenues of research. Three noted examples, in chronological order, are (1) the initial dissections of bacterioplankton communities in the Atlantic (Giovannoni et al., 1990) and Pacific (Schmidt et al., 1991) Oceans, (2) the discovery of archaeoplankton (DeLong, 1992; DeLong et al., 1994), and (3) the discovery of dominant populations of iron- and sulfur-oxidizing bacteria at hydrothermal vents (Moyer et al., 1994; 1995). This approach has now become widespread and is used in marine microbiology to apply phylogenetic analysis to establish evolutionary relationships among organisms and to use this information as a framework for making inferences about community structure, genetic and thereby inferred organismal diversity, and (to a lesser degree) to infer physiological adaptation when applicable. This approach is possible due to the detailed theory of evolutionary relationships among the domains Bacteria, Archaea and Eucarya that has emerged from comparisons of ribosomal RNA 'signature' sequences (Olsen et al., 1994b; Woese, 1994). Cell component analyses provide a culture-independent means of investigating microorganisms as they occur in nature, thereby eliminating the necessity for individual taxon cultivation (Amann et al., 1995; Ward et al., 1992). While several types of cell components are informative, SSU rDNAs (genes coding for small subunit ribosomal RNA) offer a quality and quantity of information which make them one of the most useful macromolecular descriptors of microorganisms (Ward et al., 1992). Each SSU rDNA contains both highly conserved regions which are found among all living organisms, as well as diagnostic variable regions unique to a particular population or a closely related group. SSU rDNAs are widely used as informative biomarkers for the following reasons: (1) they are essential components of the protein synthesis machinery and therefore are ubiquitously distributed and functionally conserved in all organisms; (2) they lack the interspecies horizontal gene transfer found with many prokaryotic genes; (3) they are readily isolated and identified; and (4) they 376
contain diagnostic variable regions interspersed among highly conserved regions of primary and secondary structure, permitting phylogenetic comparisons to be inferred over a broad range of evolutionary distance (Moyer et al., 1998). As a result of these studies, we are now beginning to recognize the incredible extent of diversity within the microbial world (Amann et al., 1995; Head el al., 1998; Hugenholtz el al., 1998; Ward et al., 1998). These features make SSU rDNAs particularly useful for studies of microbial ecology, where a potentially broad and unknown level of diversity of microorganisms is likely to exist. Currently, over 16 000 aligned and 30 000 unaligned SSU rRNA prokaryotic sequences have been made available for comparison by the Ribosomal Database Project II, release 8.0 (Maidak ef al., 2000), which provides these data in a phylogenetically organized format. This type of approach allows for the autecology (i.e. individual taxa) of microorganisms to be studied whether or not they can be cultivated. In addition, the phylogenetically described taxa or 'phylotypes' can be placed in a synecology context (i.e. whole community or group level) through the examination of SSU rRNA clone libraries generated from a microbial community. Depending upon the specific hypotheses to be tested, the experimental design based on molecular biological techniques can yield information regarding both autecology and synecology, in terms of community structure and phylogenetic diversity and is analogous to taking a census of a community and estimating a roadmap of evolutionary relationships for individual populations contained within. Figure 19.1 shows the dependence of environmental sample analysis with a sequence database (e.g. the Ribosomal Database Project or RDP).
METHODOLOGY FORTHE GENERATION A N D A N A L Y S I S OF SSU r D N A C L O N E LIBRARIES G e n o m i c D N A extraction and isolation The first and foremost consideration is which type of nucleic acids will be efficiently extracted from environmental samples, DNA or RNA. Once group-specific oligonucleotide probes have been constructed, and the goal is to assess to most physiologically robust components within a microbial community, then rRNA can be efficiently extracted using hydroxyapatite columns as described by Buckley et al. (1998). However, more often the generation of a clone library is needed when novel microbial communities are to be analyzed with the goal of examining microbial community structure. This requires the direct extraction of genomic DNA (gDNA) from an environmental sample. We currently use the UltraClean 'Soil' DNA Isolation kit from MoBio Laboratories, which when extracting -0.25 to 0.5 gram microbial mat samples yields approximately 5.0 to 50 lag gDNA per gram sample (wet weight). This gDNA is consistently >_10kilobases in length when gently vortexed or by using a bead beater at the lowest possible speed. This method is logistically simple and consistently
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Figure 19.1. Flowchart describing dependency of experimental design for environmental sample analysis with sequence database, while maintaining the ultimate goal of determining community structure and phylogenetic diversity.
produces purified gDNA that is able to function as substrate in restriction digests as well as template for PCR. For every sample that is processed, the concentration, purity and size are checked by spectrophotometry (i.e. 260/280 n m ratios) and by 1% gel electrophoresis against a )v-HindIII DNA standard. The residual sample debris (post-extracted) is stored at -20°C and later examined by acridine orange staining with epifluorescence microscopy to confirm cellular lysis efficiency.
378
Amplification of SSU rDNA: pitfalls and perks The success of any PCR depends largely upon the stringency of primers binding to their target template DNA during the hybridization phase. This stringency is impacted by two major factors, (1) the temperature of annealing, and (2) the concentration of free M g ions. Taq polymerase is inactive in the absence of Mg * and, with an excess, the polymerase has a greatly reduced fidelity that may increase the level of non-specific amplification. Another consideration involving a successful 'community' SSU rDNA PCR is the complexity of the template gDNA. Because multitemplate PCR is used to generate SSU rDNA clone libraries, the possibility for bias can arise, skewing the template-to-amplicon ratio. Two classes of processes have been proposed based on the theoretical modeling of PCR: (1) PCR selection and (2) PCR drift (Wagner et al., 1994). Considerable reduction in these biases has been demonstrated for SSU rDNA by using high template concentrations, performing fewer cycles, and mixing replicate reaction preparations as recommended by Polz and Cavanaugh (1998). An additional consideration is that template gDNA must be free of any RNA, otherwise single-stranded rRNA will duplex with coding strand rDNA templates thereby causing additional multitemplate bias (personal communication, Thomas Schmidt). Finally, in order to reduce tlle possibility for preferential hybridization of degenerate primers, we design and synthesize our oligonucleotides with purine and pyrimidine analogs, dK and dP, respectively (Glen Research) and with inosine where appropriate so as to minimize primer degeneracy. Primers are also synthesized with a 5' phosphalink amidite (Applied Biosystems) to facilitate ligation reactions.
Multitemplate gDNA PCR: mixtures and conditions First Master mix:
10x PCR buffer (lx final) MgC1 (2.5 m M final) 50 laM oligo primers (1 idM final for each) 2.5 mM dNTPs (200 ~M of each dNTP final) Best sterile water to 50 gl per reaction 10 mg ml ' BSA (200 ng pl ' final) 5 units Ampli-Taq Gold per reaction (Applied Biosystems) 25 mM
Second Master mix:
Combine the following master mix components for a minimum of five PCR reactions and a negative control for each SSU rDNA library to be constructed. Final volume for each reaction is 50 lal. Aliquant first master mix to each reaction tube inside a laminar flow hood using aerosol resistant pipette tips. UV irradiate for 5 to 10 rain. Then add second master mix and finally add 100 to 500 ng gDNA per reaction. No template gDNA is placed into negative control. Reaction mixtures are sealed and incubated in a thermal cycler (e.g. GeneAmp 9700; Applied Biosystems) as follows: 'hot start' at 95°C for 8 rain, 25 to 30 cycles of 94°C for I min, annealing at 55 to 60°C for 1.5 rain, with extension at 72°C for 3 min, then a final 7 min
379
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extension at 72°C, followed by a 4°C hold. Amplification products are assayed for size by 1% gel electrophoresis against a 1 kb-ladder DNA standard. Only reactions yielding no amplification of negative controls are used. Ensuing ligation step must be completed within 24 h to insure 'A' overhangs are not degraded.
Ligation, transformation and screening of SSU r D N A clones For the construction of SSU rDNA clone libraries, five independent amplification reactions from each initial sample are pooled and then quantified by spectrophotometry. This mixture is then ligated into the pTA cloning vector and transformed using the manufacturer's protocol (Clontech). Clones are screened by c~-complementation using X-gal and IPTG (-1 mg per plate each) as the substrate on LB agar plates containing 100 mg ml' ampicillin. Each putative positive clone is then selected and additionally screened by PCR using primers binding near the pTA cloning site (i.e. M13F and M13R) to determine the relative size of the insert sequence.
Putative positive screening PCR Master mix:
10x PCR buffer containing NP-40 a n d / o r TritonX-100 (lx final) 25 mM MgC1 (2.5 mM final) 5012M oligo primers (0.51aM final of both M13F and M13R) 2.5 IIIM dNTPs (250 12Mof each dNTP final) Best sterile water to 20 121per reaction 10 mg ml ~BSA (200 ng t211 final) 2 units Taq polymerase
Combine these master mix components and aliquant to each reaction tube to a final volume of 20 121inside a laminar flow hood using aerosol resistant pipette tips. A small amount of cloned cells from each white colony is then added to corresponding reactions with a sterile toothpick. The mixtures are then incubated using the previous protocol described for amplification of SSU rDNA from gDNA, except that one pre-incubation for 10 min at 94°C (to lyse the cells and inactivate any nucleases) is substituted for the 8 min 'hot start' step. Negative controls exhibiting no amplification products are required for each series of screening reactions. Amplification products are then separated and visualized on a 1% agarose gel against a 1 kb-ladder DNA standard. Clones containing correctly sized inserts are grown overnight at 37°C in N10 ml LB broth with ampicillin (100mg ml ') and are vigorously shaken. A l ml subsample of each overnight broth is aseptically transferred to a cryovial containing 0.5 ml of sterile 80% glycerol and then quick frozen and stored at -80°C. The remaining broth is used to isolate and purify plasmids using a Qiaprep spin plasmid kit according to the manufacturers protocol (Qiagen), with
380
the final plasmid elution in 100 I~1 of 0.1× Tris buffer (1.0 mM Tris-HC1, 0.1 mM EDTA, pH 8.0) and stored at -20°C.
Amplified _ribosomal D N A restriction analysis or A R D R A The ARDRA approach allows for the cataloging (based on restriction data) of SSU rDNA sequences or operational taxonomic units (OTUs) contained within a clone library thereby estimating the dominant microbial taxa contained within the sampled microbial community. The level of discrimination using four tetrameric restriction enzymes (i.e. the double-double digest) has been shown to differentiate among known SSU rDNA sequences (i.e. phylotypes) that have >98% sequence similarity (Moyer et al., 1995) and has also been found to distinguish among >99% of the bacterial taxa present within a modeled dataset of maximized diversity (Moyer et al., 1996). As ARDRA is potentially sensitive to the orientation of the cloned insert, SSU rDNA sequences are amplified from plasmid templates using oligonucleotide primers specific to proximal flanking vector sequences of the pTA plasmid. The following primers have been designed to hybridize adjacent to the pTA cloning site and are used to generate templates for the restriction digest: (5'-ACGGCCGCCAGTGTGCTG) in the forward orientation and (5'-GTGTGATGGATATCTGCA) in the reverse.
A R D R A template PCR Master mix:
10x PCR buffer (lx final) 25 mM MgC1 (2.5 mM final) 50 laM oligo primers (0.5 lJM final for both) 2.5 i'nM dNTPs (200 12Mof each dNTP final) Best sterile water to 50 lal per reaction 10 mg ml ' BSA (200 ng lal 1final) 5 units Taq polymerase
Combine these master mix components and aliquot to each reaction tube to a final volume of 50 Ill inside a laminar flow hood using aerosol resistant pipette tips, include -50 ng of purified plasmid to each reaction separately. Reactions are incubated for I rain at 95°C followed by 30 cycles of denaturation, annealing and extension at 94°C for 1 rain, 50°C for 1.5 min, and 72°C for 3 min respectively. This is followed by an additional extension at 72°C for 7 rain, and a 4°C hold. A 5 lJl subsample of each amplification is assayed for size and purity on a 1% agarose gel against a 1 kb-ladder DNA standard. Restriction digests of amplification products are performed in a microtiter dish format. Each of the two treatments (i.e. the double-double digest) consists of a well containing 15 lal of each amplification reaction and 15 pl of a restriction cocktail. Each restriction cocktail contains 3 lJl of 10x restriction digest buffer (e.g. NEBuffer 2) and either 10 units of both HhaI and HaelII or 10 units of both RsaI and MspI (New England Biolabs)
381
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per 15 #l. Restriction digest components are mixed in microtiter wells to a total volume of 30 ~1, sealed with a mylar sheet and incubated for 16 h at 37°C. After incubation, 6 1-11of Orange G loading buffer [15% (w/v) Ficoll Type 400 and 0.25% (w/v) Orange G dye] is added to each digestion reaction. DNA standards are prepared by mixing 20 lal of DNA Marker V (0.25 ~g ml '; Roche) and 4 I~1Orange G loading buffer. Separation of restriction fragments and DNA standards are performed by electrophoresis in a cold room at 4°C with 3.5% MetaPhor agarose (BioWhittaker Molecular Applications) gels run at 5 V cm ' for N4 h. Gels are stained with 0.5°7~ (w/v) ethidium bromide solution for 20 min, destained in tap water for 20 rain, and visualized by UV excitation. Gel images are captured using a digital gel documentation system (Figure 19.2). The cluster analysis of digitized restriction fragment patterns is carried out using the GelCompare software (version 4.0; Applied Maths). All gel images are digitally optimized and then normalized to a single DNA Marker V standard to reduce gel-to-gel restriction pattern variability. Cluster analysis is performed on the ARDRA patterns from all clones obtained from SSU rDNA libraries using unweighted pair group analysis of Pearson product-moment correlations. Restriction pattern clusters with correlation values between 70 and 80% are defined as discrete OTUs. As Pearson correlation coefficients are sensitive to band intensity as well as size, threshold levels must be empirically determined depending upon the type of gel documentation system used and by subjective visual examination of corresponding restriction patterns for each OTU (Figure 19.3). This process allows for an estimate of the number of representative SSU rDNA clones per OTU contained within a clone library (Heyndrickx et al., 1996).
+
+
Figure 19.2. ARDRA gel mosaic image showing double-double digest treatments in top and bottom lanes. Lanes 1, 12, 21 and 32 (designated by .) have DNA Marker V as standard, remaining lanes represent individual SSU rDNA clones. 382
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Figure 19.3. UPGMA cluster analysis of digitized and normalized ARDRA patterns indicating OTUs. Open bars on right indicate data region used in analysis which corresponds to size range of DNA standard for both treatment I and 2. OTU groupings are indicated by horizontal bars on bottom.
Rarefaction analysis In order to estimate the OTU richness as a function of diversity, the rarefaction technique is used. This is a deterministic transform of OTU abundance data. Rarefaction has the feature that it allows for the comparison of diversity from clone libraries of unequal sample size and estimates the number of phylotypes (E) in a random sample of n clones samples without replacement from a finite parent collection of N clones, where n, is the number of clones of the ith phylotype (Tipper, 1979). Rarefaction is described by the following equation:
E~=
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Rarefaction analysis with corresponding standard deviations are performed for each clone library with Matlab software (Mathworks; Moyer et al., 1998) using the algorithm developed by Simberloff (1978). A comparative example of rarified data from samples of various habitats is demonstrated in Figure 19.4.
383
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Number of clones screened Figure 19.4. Rarefaction curves as indicators of bacterial community diversity from four different habitats. Soil communities are most diverse, lake bacterioplankton community is intermediate, and hydrothermal vent microbial mat community is least diverse. All four communities were analyzed using ARDRA with the double-double digest as the basis for operational taxonomic unit (OTU) classification (Tiedje et al., 1997).
METHODOLOGY FORTHE GENERATION A N D P H Y L O G E N E T I C ANALYSIS OF SSU r D N A SEQUENCES SSU rDNA
sequencing Representative SSU r D N A clones from OTUs containing three or m o r e clones are generally the p r i m a r y targets for sequencing. The m o s t c o m m o n a p p r o a c h currently available is to use a BigDye Terminator Cycle Sequencing Kit, which uses fluorescently labeled d i d e o x y - t e r m i n a t o r s via cycle sequencing (Applied Biosystems) in conjunction with an a u t o m a t e d
384
DNA sequencer (e.g. Model 310 or 377). SSU rDNA templates used for sequencing can be generated from purified plasmids using M13F and M13R primers and PCR conditions identical to those for ARDRA analysis. Amplification products from sequencing PCR reactions are pooled and purified by size exclusion using Microcon 50 filters (Millipore) prior to sequencing. Oligonucleotides used as primers internal to the archaeal and bacterial SSU rDNA are as previously described (Lane, 1991; Moyer et al., 1998). The process of transforming raw sequence data files output by automated sequencing to contiguous SSU rDNA sequences for phylogenefic analysis is performed using the software program GeneTool with the assembly editor function (BioTools). Many programs are available that perform a similar task, however, GeneTool has been found to be extremely efficient and easy for novices to use for the purpose of 'contig' file generation and data quality control. All data should optimally be sequenced in both directions to minimize the possibility for the introduction of errors into the database.
Phylogenetic analysis: preliminary steps The first step in a successful and descriptive phylogenetic analysis is the proper alignment of SSU rDNA sequences with a collection of similar and perhaps not so similar aligned sequences from an existing database so that a hierarchical context based on molecular evolution may be inferred. This is where the Ribosomal Database Project II (RDP) functions as an invaluable resource and starting point. The RDP is an internet-accessed database (www.cme.msu.edu/RDP) that supplies phylogenetically ordered sequence alignments (their major contribution), previously constructed phylogenetic trees, ribosomal secondary structures, and distributes various software programs for constructing, analyzing, and viewing alignments and trees (Maidak et al., 2000). The usual strategy begins with a similarity search using a newly generated SSU rDNA sequence to query the database for sequences that are the most similar. This can be accomplished directly through the RDP using the SEQUENCE MATCH utility and also by using a basic BLAST search for the latest Genbank accessions (www.ncbi.nlm.nih.gov). This approach achieves two tasks, first to find if any identical or closely related sequences exist in the database and second to ascertain the level of dissimilarity between a potentially novel sequence and any previously recorded phylogenetic groups. Both of these searching functions are based on estimating S,~,values and cannot be used to infer in-depth phylogenetic relationships. Another consideration regarding multitemplate PCR of SSU rDNAs is the potential generation of non-extant chimeras and thus artifactual sequences leading to the erroneous description of nonexistent microorganisms. At this point, sequences should be submitted to detect possible chimeric artefacts using the nearest-neighbor based CHECK CHIMERA function online at RDP (Robinson-Cox ct al., 1995) a n d / o r the k-tuple
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matching method of mglobalCH1 available at www-hto.usc.edu/ software/mglobalCHI (Komatsoulis and Waterman, 1997). Chimeras are certainly not a rarity and every sequence must be thoroughly tested, including a complete secondary structure analysis looking for noncompensatory base changes. Chimeras have been found to occur at -5% in multitemplate clone libraries even under the most stringent of PCR conditions. However, an advantage of the ARDRA approach is that no chimera sequence has occurred more than once within any OTU detected from any single clone library. Once this stage has been completed, then the initial choices for comparative microbial sequences used in the phylogenetic analysis can be made. The next phase is by far the most critical step in an accurate phylogenetic analysis regardless of the algorithm used to model evolutionary distance. Phylogenetic analysis is restricted to the comparison of highly to moderately conserved nucleotide positions that are unambiguously alignable in all sequences to be examined. The basic assumption is that these data then represent homologous positions of common ancestry. This step involves the alignment of novel sequences to previously aligned sequences, which again can be obtained from the RDP. One must realize that although the alignment of sequences is relatively simple among closely related taxa, it can be very difficult as the sequences become more divergent. Multiple sequence alignments can be constructed with programs such as the Genetic Data Environment (GDE) distributed by RDP or with the graphically oriented 'ARB: a software environment for sequence data' (www.biol.chemie.tu-muenchen.de) which links sequence data files to a dendrogram hierarchy (Strunk et al., 1998). The ARB package has the added advantage of an automated aligner function. However, in either case, this process weighs heavily upon secondary structure considerations and alignments must be checked against known secondary structures, as all rRNA molecules regardless of ancestry share a common core of secondary structure. Generally, this process is achieved by the construction of a 'mask' or row of l's and O's allowing the phylogenetic algorithm to process specific columns of data from the alignment file. Since data removal means information loss, it is advantageous to analyze each dataset with multiple mask variations. This potentially shows the robustness of a given tree topology and gives an estimate as to whether there is a substantial influence from the more highly variable positions. Both ARB and GDE have the capacity to use weighted masks with multiple sequence alignments.
Phylogenetic analysis: which
algorithm should I use?
There are basically three approaches used for the reconstruction of phylogenetic trees: distance matrix, maximum parsimony, and maximum likelihood methods. These algorithms are based on evolutionary models with different criteria for estimating evolutionary distance and maximizing the congruency of tree topologies (Ludwig et al., 1998). Assumptions common among each of these approaches are: (1) each character is ew)lving
386
independently; (2) nucleotide changes are primarily neutral; (3) comparisons are among orthologous genes; and (4) positional homology has been inferred correctly. Distance matrix methods revolve around a two-step approach where first a matrix of pairwise distance values is calculated based on various nucleotide substitution formulas (i.e. the Jukes and Cantor one-parameter or Kimura two-parameter models). Then after the distance matrix is calculated, binary sequence differences are transformed into a tree using a clustering algorithm such as the neighbor-joining or DeSoete methods. This approach is advantageous when many taxa are compared and highthroughput tree building is necessary as it is computationally the least expensive. The disadvantages are that sequence data is converted into distance values, thereby reducing some phylogenetic information. Overall, distance matrix methods represent a compromise, but are especially useful for initial phylogenetic screening or when taxa for diverse and yet established lineages are compared (Figure 19.5). Both the ARB and GDE (with the inclusive PHYLIP software) packages are able to produce distance matrices and generate trees from distance data. The remaining two approaches are both character-based methods where the aligned sequence data (i.e. individual nucleotide positions) are used directly by the respective algorithm. Maximum parsimony is popular due to its logically simple and truly cladistic model known as Ocham's Razor, where the simplest solution is decidedly the best solution assuming that homoplasy (i.e. parallelism or convergence) is minimal. This is where the selected tree(s) has/have the shortest overall tree length and is supported by the largest number of synapomorphies (i.e. shared and derived character sites). The disadvantages are that maximum parsicnony relies heavily upon synapomorphies (i.e. much information is lost) and a single best-fit tree may not necessarily be found. Also, it requires a greater computational capacity than any of the distance matrix methods. ARB and the new PAUP* (Sinauer) are examples of software packages which allow both the estimation of branch lengths as well as the generation of trees according to maximum parsimony. The maximum likelihood approach for tree reconstruction is the most sophisticated and robust of the three methods, and allows for the inequality of transition and transversion rates. This statistically motivated approach calculates the tree for which the observed data are most probable, using a given nucleotide substitution model. The algorithm itself functions as a two-step process where first it defines the tree topology and then optimizes the branch lengths on that particular topology (Felsenstein, 1981). The big advantage is that this method uses all of the character data and as such looks at every possible scenario of evolutionary change at each nucleotide position. The primary disadvantage is that due to the tremendous number of calculations it is by far the most computationally intensive. However, using the enhanced version (i.e. fastDNAml) which significantly improves computational performance (Olsen et al., 1994a) and with the advent of modern computer technology, this has become much less of a burden and enabled phylogenetic tree reconstruction with _>25 taxa with a Sun workstation (Figure 19.6). Trees 387
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are constructed using jumbled orders for the addition of taxa and allowing for the global swapping of branches. Using these parameters, the search for an optimal tree is repeated until the best log likelihood score is reached in at least three i n d e p e n d e n t searches. The fastDNAml program is also distributed by the RDE In order to further test the confidence of branching orders, resampling techniques such as bootstrapping can be used in conjunction with any of the phylogenetic approaches so that node reproducibility and robustness can be determined (Felsenstein, 1985). Bootstrap values are assigned to 388
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Figure 19.6. Phylogenetic tree demonstrating the relationships of the PV-1 and ES-1 cultured isolate phylotypes, which are included in Guaymas Vent Bacteria (GVB OTU 1) and Pele's Vents Bacteria (PVB OTU 1) lineage, with other ?Proh,obacteria and additional representative iron- and sulfur-oxidizers, as determined by m a x i m u m likelihood analysis of SSU rDNA sequences. Numbers at nodes represent bootstrap values (percent) for that node (based on 200 bootstrap resamplings). An outgroup is represented by Arlhrot~ach'r slobifor,Hs. The scale bar represents 0.10 fixed mutations per nucleotide position. Bootstrap values are shown for frequencies at or above a threshold of 50% (Emerson and Moyer, 1997; unpublished data).
each internal node of a tree, indicating the percentage of the time that a subtree defined by that respective branch appears as monophyletic. When used with fastDNAml, generally a threshold of >_50c/~ is used and bootstrapping occurs _>100 times again with a jumbled addition of taxa and the search for each optimal tree is repeated until the best log likelihood score is reached in at least two i n d e p e n d e n t searches (Figure 19.6). The collection of bootstrapped trees is compiled using the consensus tree function in either the GDE (with the inclusive PHYLIP software) or PAUP* software packages in order to calculate bootstrap values. For a comprehensive
389
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review of the methods used in phylogenetic analysis, including an indepth description of the mathematical modeling and theory, see Swofford et al. (1996).
CONCLUDING
REMARKS
This chapter describes an avenue for the application of m o d e r n molecular biological techniques to marine microbiology. Many promising molecular-based applications are also viable alternatives such as fluorescent in situ hydridization (FISH) of group-specific oligonucleotide probes (Amann et al., 1995) or the high-throughput m e t h o d of terminal restriction fragment length p o l y m o r p h i s m (T-RFLP) used to track specific populations through space and time (Marsh et al., 2000). However, as s h o w n in Figure 19.1, environmental sample analysis remains d e p e n d e n t upon the available database of k n o w n (and aligned) sequences. This, coupled with the observation that >>1% of physiologically defined microorganisms found in culture collections have been detected in environmental samples, points to the efficacy of the clone library approach coupled with the phylogenetic analysis of SSU rDNA sequences w h e n attempting to understand the microbial c o m m u n i t y structure and diversity from marine habitats.
References Amann, R. I., Ludwig, W. and Schleifer, K. H. (1995). Phylogenetic identificafion and ill situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59, 143-169. Buckley, D. H., Graber, J. R. and Schmidt, T. M. (1998). Phylogenetic analysis of nonthermophilic members of the kingdom Crenarchaeotn and their diversity and abundance in soils. Appl. Environ. Microbiol. 64, 4333 4339. DeLong, E. E (1992). Archaea in coastal marine environments. Proc. Natl. Acad. Sci. USA 89, 5685 5689. DeLong, E. E, Wu, K. Y., Prezelin, B. B. and Jovine, R. V. M. (1994). High abundance of Archaea in Antarctic marine picoplankton. Nature 371, 695-697. Emerson, D. and Moyer, C. L. (1997). Isolation and characterization of novel ironoxidizing bacteria that grow at circumneutral pH. Appl. El~viron. Microbiol. 63, 4784 4792. Felsenstein, J. (1981). Evolutionary trees from DNA sequences: a maximum likelihood approach. J. Mol. Evol. 17, 368-376. Felsenstein, J. (1985). Confidence Iimi¢s on phykGenies: an approach using bootstrap. Evolution 39, 783-791. Giovannoni, S. J., Britschgi, T. B., Moyer, C. L. and Field, K. G. (1990). Genetic diversity in Sargasso Sea bacterioplankton. Nature 345, 60-63. Head, I. M., Saunders, J. R. and Pickup, R. W. (1998). Microbial evolution, diversity, and ecology: A decade of ribosomal RNA analysis of uncultivated microorganisms. Microbial. Ecol. 35, 1-21. Heyndrickx, M., Vauterin, L., Vondamme, I~, Kersters, K. and De Vos, P. (1996). Applicability of combined amplified ribosomal DNA restriction analysis (ARDRA) patterns in bacterial phylogeny and taxonomy. ]. Microbiol. Meth. 26, 247-259.
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Hugenholtz, R, Goebel, B. M. and Pace, N. R. (1998). h'npact of culture-independent studies on the emerging phylogenetic view of bacterial diversity I. Bacteriol. 180, 4765-4774. Karl, D. M. (1986). Determination of in situ microbial biomass, viability, metabolism, and growth. In: Bacteria i, Nature (J. S. Poindexter and E. R. Leadbetter, Eds), Vol. 2, pp. 85-176. Plenum Press, New York. Komatsoulis, G. A. and Waterman,. M. S. (1997). A new computaional method for detection of dlimeric 16S rRNA artifacts generated by PCR amplification from mixed bacterial populations. Appl. EnviroH. Microbiol. 63, 2338-2346. Lane, D. J. (1991). 16S/23S rRNA sequencing. In: Nucleic Acid Techniques in Bacterial Systematics (E. Stackebrandt and M. Goodfellow, Eds.), pp. 115-175. John Wiley & Sons, Ltd., London, England. Ludwig, W, Strunk, O., Klugbauer, S., Klugbauer, N., Weizenegger, M., Neumaier, J., Bachleitner, M. and Schleifer, K. H. (1998). Bacterial phylogeny based on comparative sequence analysis. Eh'ctrophorcsis 19, 554-568. Maidak, B. L., Cole, J. R., Lilburn, G., Parker, C. T., Saxman, R R., Stredwick, J. M., Garrity, G. M., Li, B., Olsen, G. J., Pramanik, S., Schmidt, T. M. and Tiedje, J. M. (2000). The RDP (Ribosomal Database Project) continues. Nucl. Acids Res. 28, 173-174. Marsh, T. L., Saxman, P., Cole, J. and Tiedje, J. (2000). Terminal restriction fragment length polymorphism analysis program, a web-based research tool for microbial analysis. Appl. Enviro,. Microbiol. 66, 3616 3620. Morita, R. Y. (1986). Autecological studies and marine ecosystems. In: Microbial Autecology: A Method for Enviromm'ntal Stm[ies (R. L. Tare, Ed.), pp. 147-181. John Wiley & Sons, Ltd., London, England. Morita, R. Y., and Moyer, C. L. (2000). Biodiversity of psychrophiles. In: E,cyclopedia ~ff Biodiversity, (S. A. Levin, R. Colwell, G. Daily, J. Lubchenco, H. A. Mooney, E.-D. Schulze, G. D. Tilman, Eds). In press. Academic Press, San Diego. Moyer, C. L., Dobbs, E C. and Karl, D. M. (1994). Estimation of diversity and community structure through restriction fragment length polymorphism distribution analysis of bacterial 16S rRNA genes from a microbial mat at an active, hydrothermal vent system, Loihi Seamount, Hawaii. ApFI. Enviro,. Microbiol. 60, 871-879. Moyer, C. L., Dobbs, E C. and Karl, D. M. (1995). Phylogenetic diversity of the bacterial community from a microbial mat at an active, hydrothermal vent system, Loihi Seamount, Hawaii. Appl. E,viroH. Microbiol. 61, 1555-1562. Moyer, C. L., Tiedje, J. M., Dobbs, E C. and Karl, D. M. (1996). A computer-simulated restriction fragment length polymorphism analysis of bacterial small subunit rRNA genes: Efficacy of selected tetrameric restriction enzymes for studies of microbial diversity in nature. Appl. Enviro,. Microbiol. 62, 2501-2507. Moyer, C. L., Tiedje, J. M., Dobbs, F. C. and Karl, D. M. (1998). Diversity of deepsea hydrothermal vent Archaea. Deep-Sea Res. 11.45, 303-317. Olsen, G. J., Lane, D. J., Giovannoni, S. J. and Pace, N. R. (1986). Microbial ecology and evolution: a ribosomal RNA approach. A , , . Roy. Microbiol. 40, 337-365. Olsen, G. J., Matsuda, H., Hagstrom, R. and Overbeek, R. (1994a). fastDNAml: a tool for construction of phyk)genetic trees of DNA sequences using maximum likelihood. CABIOS 10, 41-48. Olsen, G. J., Woese, C. R. and Overbeek, R. (1994b). The winds of (evolutionary) change: breathing new life into microbiology. Microbiol. Rev. 176, 1 6. Pace, N. R., Stahl, D. A., Lane, D. J. and Olsen, G. J. (1986). The analysis of natural microbial populations by ribosomal RNA sequences. Adv. Microb. Ecol. 9, 1-55. Polz, M. F. and Cavanaugh, C. M. (1998). Bias in template-to-product ratios in multitemplate PCR. Appl. EnviroiT. Microbiol. 64, 3724-3730.
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Robison-Cox, J. E, Bateson, M. M. and Ward, D. M. (1995). Evaluation of nearestneighbor methods for detection of chimeric small-subunit rRNA sequences. Appl. Environ. Microbiol. 61, 1240-1245. Schmidt, T. M., DeLong, E. E and Pace, N. R. (1991). Analysis of a marine picoplankton community by 16S rRNA gene cloning and sequencing. ]. Bacteriol. 173, 4371-4378. Simberloff, D. (1978). The use of rarefaction and related methods in ecology, in:
Biological Data in Water Pollution Assessment: Quantitative and Statistical Analyses (K. L. Dickson, J. Cairns, Jr. and R. J. Livingston, Eds), pp. 150-165. ASTM STP 652. Strunk, O., Ludwig, W., Gross, O., Reichel, B., Stuckmann, N., May, M., Nonhoff, B., Lenke, M., Ginhart, T., Vilbig, A. and Westram, R. (1998). ARB: A Software Environment for Sequence Data. Department of Microbiology, Technical University of Munich, Munich, Germany. Swofford, D. L., Olsen, G. J., Waddell, P. J. and Hillis, D. M. (1996). Phylogenetic inference. In: Molecular Systematics 2nd edn (D. M. Hillis, C. Morffz, B. K. Mabel, Eds), pp. 407-514. Sinauer, Sunderland, MA. Tiedje, J. M., Zhou, J.-Z., N~isslein, K., Moyer, C. L. and Fulthorpe, R. R. (1997). Extent and patterns of soil microbial diversity. In: Progress in Microbial Ecology: Ptvceedings of the 7th International Symposium on Microbial Ecology/(M. T. Martins, M. I. Z. Sato, J. M. Tiedje, L. C. N. Hagler, J. D6bereiner and P. S. Sanchez, Eds.), pp. 35-41. Brazilian Society for Microbiology, S~o Paulo, Brazil. Tipper, J. C. (1979). Rarefaction and rarefiction - - the use and abuse of a method in paleoecology. Paleobiology 5, 423-434. Wagner, A., Blackstone, N., Cartwright, P., Dick, M., Misof, B., Snow, P., Wagner, G. 1{, Bartels, J., Murtha, M. and Pendleton, J. (1994). Surveys of gene families using polymerase chain reaction: PCR selection and PCR drift. Syst. Biol. 43, 250-261. Ward, D. M., Bateson, M. M., Weller, R. and Ruff-Roberts, A. L. (1992). Ribosomal RNA analysis of microorganisms as tiley occur in nature. Adv. Microb. Ecol. 12, 219-286. Ward, D. M., Ferris, M. J., Nold, S. C. and Bateson, M. M. (1998). A natural view of microbial biodiversity within hot spring cyanobacterial mat communities. Microbiol. Mol. Biol. Rev. 62, 1353-1370. Woese, C. R. (1994). There must be a prokaryote somewhere: microbiology's search for itself. Microbiol. Rev. 58, l-9. ZoBell, C. E. (1946). Marine Microbiology: A Monof~raph ou Hydrobacteriolo~y. Chronica Botanica Co. Waltham, MA.
List of suppliers Applied Biosystems
Applied Maths BVBA
850 Lincoln Centre Drive Foster City, CA 94404, USA Tel.: 650-570-6667 1-877-477-3675 Fax: 650-572-2743 Web: www.appliedbiosystems.com
Risquons- Toutstraat 38 8511 Kortrijk, Belgium Tel.: 32-56-424144 Fax: 32-56-402145 Web: www.applied-maths.com
B i g D y e Terminator Cycle S e q u e n c i n g Kit.
392
GelCompar software program.
BioTools Incorporated 420 Sun Life Place 10123 99 Street Edmonton, Alberta, Canada T5J 3H1 Tel.: 1-780-423-1133 Fax: 1-780-423-1333 Web: www.biotools.com
Mo Bio Laboratories, Inc. P.O. Box 606 Solana Beach, CA 92075, USA Tel.: 760-929-9911 1-800-606-6246 Fax: 760-929-0109 Web: www.mobio.com
GeneTool software program.
'Soil' DNA Isolation Kit.
co
BioWhittaker Molecular Applications 191 Thomaston Street Rockland, MD 04841, USA Tel.: 207-594-3400 1-800-341-1574 Fax: 207-594-3426 Web: www.bmaproducts.com
N e w England Biolabs 32 Tozer Road Beverly, MA 01915, USA Tel.: 1-800-632-5227 Fax: 1-800-632-7440 Web: ~UWW.tleb.cotll
Source of Tetrameric Endonucleases.
MetaPhor agarose for high resolution separation of small DNA fragments. Clontech Laboratories, Inc. 1020 East Meadow Circle Palo Alto, CA 94303, USA Tel.: 650-424-8222 1-800-662-2566 Fax: 650-424-1064 Web: www.clolltec]l.coltl
Qiagen Inc. - USA 28159 Avenzle Stanford Valencia, CA 91355, USA Tel.: 1-800-426-8157 Fax: 1-800-718-2056 Web: www.qiagen.com
QIAprep Spin Plasmid Minprep Kit.
AdvanTAge PCR Cloning Kit. Glen Research 22825 Davis Drive Sterling, VA 20164, USA Tel.: 703-437-6191 1-800-327-4536 Fax: 703-435-9774 W~b: WWW. ~[ellFCS.COttl
Roche Molecular Biochemicals 9115 Hague Road P.O. Box 50414 Indianapolis, IN 46250, USA Tel.: 1-800-428-5433 Fax: 1-800-428-2883 Web: biochem.roche.conl
Supplier of low molecular weight DNA standard Marker V.
dK and dP nucleotide analogs. Millipore Corp. 80 Ashby Road Bedford, MA 01730, USA Tel.: 1-800-645-5476 Fax: 1-781-533-3110 Web: www.millipore.com
Microcon 50 ultrafiltration units.
393
Sinauer Associates, Inc. P.O. Box 407 23 Plumtree Road Sunderland, MA 01375, USA Tel.: 413-549-4300 Fax: 413-549-1118 Web: wu_rLi.Lsillaller.cotll
PAUP* 4.0 (beta version) software programs.
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The MathWorks, Inc. 3 Apple Hill Drive Natick, MA 01760, USA Tel.: 508-647-7000 Fax: 508-647-7001 Web: www.matl~works.com Matlab software program used with 'Rarefier' program.
394
20 Gene Expression by mRNA Analysis John H Paul Department of Marine Science, University of South Florida, St Petersburg, FL 33 70 I, USA Z
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INTRODUCTION Contemporary studies in marine microbial ecology have attempted to address the fundamental question of 'Who's out there?' from a standpoint of molecular diversity of the populations present (see Chapter 19 by Moyer). An enormous amount of sequence information has been catalogued about marine microbial communities. To relate genetic information to the activities of microbial populations requires a knowledge of gene expression (MacGregor, 1999). Gene expression encompasses transcription, translation, post-translational modification, and production of a functional gene product. A first approach to this problem has been the detection of gene expression as transcriptional activity, or the detection of specific target mRNAs (Pichard and Paul, 1991; Pichard et al., 1993). Although this approach does not provide information on a functional gene product, or phenotype, it does yield information on how specific genes are activated in environmental populations. Thus, questions concerning molecular regulation of enzymes that perform vital functions in the environment (nutrient cycling, bioremediation, and biogeochemical processes) can now be answered. The transcriptional activity approach has been taken to understand the pathways of carbon and nitrogen cycling in the environment. For example, Wyman et al. (1996) demonstrated for the first time clear METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0-12 521530-4
Copyright © 2001 Academic Press Lid All rights of reproduction in any form reserved
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patterns of diel regulation in nifH (which encodes for the iron-containing subunit of nitrogenase) in natural populations of Trichodesmium thiebaultii. In this case, levels of nifH transcripts were tightly linked to nitrogenase activity, which peaked early in the daylight hours (800 to 1200 h). This work definitively showed that there was no temporal separation between nitrogen fixation, an anaerobic process, and oxygenic photosynthesis, as has been observed for Synechococcus RF-1 (Chow and Tabita, 1994). Diel rhythms in carbon fxation and glutamine synthetase activity were noted in natural populations of Synechococcus populations (Wyman, 1999), with rbcL peaking in the morning, and glnA maxima at 1400 h. Glade-specific, diel rhythms in rbcL expression have been observed in natural populations of phytoplankton off Cape Hatteras and in the Gulf of Mexico (Paul et al., 1999b), with cyanobacterial rbcL expression occurring in the morning, and diatom-like rbcL transcripts appearing in the afternoon or early evening. The role of iron in nitrogen regulation in Synechococcus has been investigated using mRNA analysis. Both Fe and nitrate seemed to cause induction of niR (nitrate reductase) gene transcripts (Sadhegi, 1998). The induction of psbA2 and psbA3 by UV light, which encode for components of Photosystem II (PSII), appears to be a defense response against UV damage of PSII in Synechocystis PCC6803 (Mate et al., 1998). The heat shock protein hspA mRNA was shown to be produced in response to exposure to heat in the thermophilic cyanobacterium Synechococcus vulcanus (Roy and Nakamoto, 1998). Thus, mRNA analysis has enabled determination of the response of natural populations to perturbations in environmental conditions. As well as photoautotrophs, gene expression studies have been very important in the study of marine heterotrophic bacteria, particularly the extremophiles, in terms of understanding the response to pressure (Bartlett and Welch, 1995), and low and high temperature (Hamamoto et al., 1995; Robinson et al., 1994). This approach has also been taken to investigate the control of chitin degradation in the marine bacterium Pseudoaltelvmorlas strain $91 (Techkarijanauk and Goodman, 1999). The capability to detect gene expression by mRNA analysis has enabled the real-time detection of expression of genes involved in bioremediation as well. Naphthalene dioxygenase activity has been detected in a coal tar contaminated site, and analysis of clones obtained by RT-PCR indicated the presence of novel genotypes involved in this process (Wilson et al., 1999). As more interest arises in the response of organisms to environmental changes, conditions, and substrate availability, the need for environmental gene expression studies continues to increase. In this paper we describe three related methods for mRNA isolation, and compare some results obtained by each.
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S T A N D A R D P R E C A U T I O N S FOR M A K I N G RNAse-FREE S O L U T I O N S A N D W O R K I N G WITH RNA RNAses are ubiquitous and autoclaving will not fully inactivate all RNAses. Therefore, it is necessary to avoid contamination of solutions and lab supplies when working with cellular RNA. It is recommended that disposable gloves be worn when working with RNA and in the preparation of reagents to be used in RNA work. Whenever possible, sterile disposable labware should be used in working with RNA. When glassware is to be used, it is important to decontaminate by washing in 0.5 N NaOH and then rinsing with deionized water. The glassware is then baked at 450°C overnight, with glassware openings covered with aluminum foil. It is important to remove all plastic from the glassware, and to treat plastic lids and sealing rings with diethyl pyrocarbonate (DEPC), followed by autoclaving (45 min) to remove RNAses. Glass beads used for bead beating are aliquoted into foil packets (10 g per packet) prior to baking. Metal spatulas for weighing out reagents should be washed with 0.5 N NaOH and rinsed with 100% ethanol and flamed. All reagents for use with RNA should be separate stocks for that purpose only and not from the general lab supplies. All dry chemicals used for making reagents should also be designated for RNA work only. If possible, a separate lab area and equipment should be set out for work with RNA. Areas and equipment can be wiped with 1% sodium dodecyl sulfate (SDS) or 0.5 N NaOH, ethanol, and DEPC-treated water successively to reduce contamination if necessary. To ensure that solutions are free of RNAse contamination they should be treated with DEPC at a concentration of 0.1~. overnight at 37°C in a fume hood (DEPC is a suspected carcinogen). Autoclave the solutions twice for 30 min or more, as this causes breakdown of DEPC to ethanol and carbon dioxide. Solutions containing primary amines (i.e. Tris) should not be used with DEPC because of potentially explosive reactions. DEPC will cause polycarbonate (such as that in filtration towers, dot blot apparatus, etc.) to become brittle and cracked. These materials should be washed with 1% SDS instead, rinsed with ethanol and then autoclaved. Biochemical reagents that cannot be autoclaved (such as nucleotides, dithiothreitol, etc.) should be made up with DEPC-treated water and sterilized by filtration. To prevent RNAse contamination of pH probes, the probe should be treated with 3% sodium peroxide (made from a 10% stock using DEPCtreated water) for 10 min in a baked glass beaker. The pH buffers and probe should be dedicated for RNA work only. The probe should be rinsed between uses with DEPC-treated DI. When amplification is the ultimate goal, it is vital that RNA extraction and isolation be performed in a different location (preferably a separate room) from amplification and detection. This prevents contamination of the extracted matter with amplicon and resultant false positive amplifications. 397
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Safety precautions when working with RNA DEPC, phenol, and chloroform are all potential carcinogens. Be certain that they are used in a recently calibrated fume hood. All RNA extraction reagents described below use guanidinium isothiocyanate which is also extremely toxic and liberates toxic gas, and should be used in a fume hood. Wear a lab coat and gloves to protect the skin. No mouth pipetting, eating, or smoking should occur in areas in which RNA extraction is to occur.
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COLLECTION OF SAMPLES FROM NATURAL POPULATIONS FOR RNA ANALYSIS A problem with collecting samples for mRNA analysis of natural phytoplankton populations is that there is often a very low biomass present, which requires concentrating cells from large volumes of water. The only recourse is to filter as rapidly as possible and to minimize nuclease degradation of RNA by DEPC-treating water samples. The method outlined below has proven successful for m a n y environments sampled, and is adapted for shipboard use.
Materials and supplies •
• • • • • •
Filtration manifold, polyvinylchloride or stainless steel, equipped with glass towers and stainless steel filtration grids (25 mm diameter). We have modified these towers by attaching funnels to the top made from the tops of polypropylene bottles (equivalent to Chlorox TM bottles). Each tower holds about 200 ml. Portable BeI-Art TM polypropylene fume hoods and exhaust fans. Each holds two manifolds and are an absolute necessity because of work with DEPC. DEPC (diethyl pyrocarbonate), 100%, Sigma Chemical Corporation. Millipore Durapore filters, autoclaved, 0.45 pm pore size, 25 mm in diameter (0.22 pm pore size results in no recovery of RNA). I gallon (3.78 I) polypropylene side-arm jugs for short-term sample storage. Dewar with liquid nitrogen. 2.0 ml screw cap microcentrifuge tubes, sterile, containing the appropriate volume of extraction reagent, depending upon the extraction method to be employed.
Procedure . The water sample is collected and transported to the filtration site as quickly as possible, preferably within minutes. For lakes and rivers, it is advised to have a mobile lab with filtration capabilities as described above. We have found it convenient to put samples in the polypropylene jugs as they are collected from Niskin bottles. 398
2. DEPC is immediately added to the sample in a fume hood (or out of doors) for a final concentration of 0.01%. This should be done in the polypropylene jugs and the jugs capped. Note If the sample is to be used for RT-PCR do not add DEPC as this might cause RNA cross-linking that will interfere with the RT reaction. 3. The samples are filtered onto the Durapore filters. We usually filter 0.5-1 1 for oligotrophic, offshore waters and 200-400 ml for estuarine waters. (This requires patience!) 4. The filters are folded in half and inserted into 2.0 ml microcentrifuge tubes that contain the extraction reagent (see below), the tubes labeled, and immediately frozen in liquid nitrogen. The samples are then stored at-80°C until analysis.
GIPS M E T H O D FOR R N A I S O L A T I O N The GIPS (Guanidinium Isothiocyanate Phenol Sarcosyl) method is based upon the method of Chomczynski and Sacchi (1987) and was commercialized into a kit by Cinna/Biotecx under the name RNAzol ~', which is no longer available. A newer product has been developed which results in the simultaneous isolation of RNA, DNA, and protein, called Tri-Reagent 1~ (Molecular Research Center), which we have not evaluated. We have found more reproducible results if we made the reagents ourselves rather than relying on commercial preparations (shelf life we estimate to be about one month). The basic method takes advantage of isolation of RNA in the presence of the strong denaturant, guanidinium isothiocyanate, along with the detergent sarcosyl, to facilitate cell lysis. The method was developed for human and animal cells and requires the addition of mechanical disruption techniques (i.e. bead-beating) to lyse procaryotic and many algal cell types (Pichard and Paul, 1991). The length of time for bead-beating should be evaluated for each application (Tim Hopkins, Biospecs, personal communication). We recommend 2 rain pulses, with ice treatment in between, up to perhaps 10 min total. The method is unique in that it employs acidified phenolic extraction which results in the accumulation of DNA at the phenol/aqueous interface, leaving pure RNA in the aqueous supernatants. Thus, of all the methods we have employed, this results in the least contamination of the sample with DNA. A clear disadvantage is the use of phenol and production of phenol/chloroform waste as well as the time required and extensive re-precipitation.
Equipment and reagents •
•
Biospec Products Mini-Bead-BeaterTM or Bead-Beater-8 TM. The Mini-BeadBeater uses 2.0 ml bead-beater cubes (one at a time) whereas the BeadBeater-8 can accommodate eight cubes at once. Bead-beater vials (2_.0ml screw cap microfuge tubes with O-rings; Biospec Products or Fisher Scientific #05-664-34).
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Glass beads (0.1-0.15mm diameter, Biospec Products, cat #11079-101), muffle-furnace treated (450 °) for 4 h. GIPS extraction reagent: to 28.4 g guanidinium isothiocyanate (Fisher, Molecular Biology Grade) add twice autoclaved, DEPC-treated deionized water (at 60-65°C) using a DEPC-treated stir bar. After this is dissolved, add 3 ml 10% sarcosyl solution (at 60-65°C) and 2 ml 0.75 M sodium citrate, pH 7.0. 10% sodium sarcosyl:add twice autoclaved, DEPC-treated deionized water to 6 g sarcosyl (N-lauryl sarcosine) to a total volume of 60 mI. Autoclave and store at room temperature. 0.75 M sodium citrate, pH 7.0.Add twice autoclaved, DEPC-treated deionizecl water (-20 ml) to 13.23 g trisodium citrate.Adjust the pH to 7.0 with a drop or two of concentrated HCI, or 8 N NaOH. Bring the final volume to 60 ml with DEPC deionized water, autoclave, and store at room temperature. 2 M sodium acetate, pH 4:Add 40 ml twice autoclaved, DEPC-treated deionized water and 35 ml glacial acetic acid to 16.4 g sodium acetate.Adjust the pH to 4 with additional acetic acid and the final volume to 100 ml with sterile, DEPC-treated deionized water. Water-saturated phenol and chloroform/isoamyl alcohol (49:1). Make these as per Sambrook et al. (1989) except use DEPC-treatecl DI in the preparation of phenol.
Procedure 1. Samples are either collected by filtration onto sterile 25 m m 0.45 btm Durapore (Millipore) filters or the samples are collected by centrifugation (cell cultures). 2. The filter is placed into a sterile 2.0 ml bead beater tube containing: (a) 0.5 g of baked glass beads (8-9 scoops of a small spatula) (b) 0.5 ml of GIPS extraction reagent (c) 50 btl of sodium acetate pH 4 (d) 0.5 ml of water-saturated phenol (e) 0.1 ml of chloroform:isoamyl alcohol (49:1). 3. The tube is placed in a Mini-Bead-Beater or a Bead-Beater-8 and processed for 2 rain. 4. The sample is placed on ice for 15 rain and then centrifuged in a microcentrifuge for 5 min to separate the aqueous and organic fractions. 5. The top (aqueous) phase is recovered to a new tube and the sample reextracted two more times by adding 0.5ml of GIPS and 50~tl of sodium acetate to the original tube containing the filter and the original phenol/chloroform, followed by steps 3 and 4 above. 6. The aqueous extracts are combined, 10 btg glycogen added, and precipitated with one volume of isopropanol for 2 h at -20°C. 7. The precipitated RNA extracts are microcentrifuged for 10 min and the RNA is taken up in 200 btl of 1 mM EDTA pH 7.0. 8. The sample is reprecipitated by addition of 0.1 vol 3 M NaC1, 10 pg glycogen, and two volumes of 95% ethanol overnight at -20°C. 9. The RNA is again collected by microcentrifugation, washed twice with one volume of ice-cold 70% ethanol, and re-suspended in 10 to 30 btl of 400
1 mM EDTA pH 7.0 (10 gl for RT-PCR, 30 gl for blotting and quantitation of mRNA). 10. The sample may then be dotted directly (see below), used in RT reactions, or stored at -80°C.
B O O M M E T H O D FOR R N A I S O L A T I O N This method is an adaptation of the guanidinium isothiocyanate extraction technique based on the method of Boom et al. (1990). The advantage of the method is that it does not require phenol but rather takes advantage of the ability of nucleic acids to bind to silica. A commercially available kit is manufactured by Organon Teknika (NucliSens TM) which works basically on the same principal. A disadvantage of the method is that unlike methods which include alcoholic precipitation, where very high recoveries are ensured, the nucleic acids may not be totally removed from the silica. A second disadvantage is that the method is not specific for isolation of RNA but extracts both DNA and RNA (see Results section). Therefore, before quantitative hybridization for transcript levels can be accomplished, it is necessary to digest with an RNAse-free DNAse.
Equipment and reagents •
Silica suspension: 60 g silica (Sigma $563 I; M W 60.08; part. Size I-S ~[m) is suspended in 500 ml DI water in a SO0 ml graduated cylinder for 2_4h. The supernatant is drawn off until 70 ml remains. An additional 500 ml of DI water is added, and the mixture is shaken to re-suspend the silica. Allow this to settle for 5 h at room temperature. Draw off the supernatant until 60 ml remains.Add 533 ~l of concentrated HCI, then add 60 pl of DEPC. Autoclave twice for 20 min, and then aliquot to 2_ml tubes and freeze at -20°C. • 0.2 M EDTA: dissolve 37.2_g of EDTA and 5 g of NaOH in 450 ml of DI water. Adjust pH to 8.0 with NaOH and adjust volume to 500 ml. Autoclave. • L2 buffer: dissolve 12.1 g of Tris-Base in 800 ml of DI water.Adjust the pH to 6.4 with concentrated HCI, and bring the volume to 1000ml with DI. Autoclave. • L2 wash buffer: to 100 ml of L2 buffer add 120g of guanidine thiocyanate (guanidinium isothiocyanate-Molecular Biology Grade, Fisher Biotech). • L6 lysis buffer: to 100 ml of L2 buffer add 22 ml of 0.2 N EDTA (pH 8.0) and 2.6 g of Triton X-100.The pH is then adjusted to 6.4, and 120 g of guanidinim isothiocyanate is added, it is often necessary to facilitate dissolution by autoclaving. • Muffle furnace-treated glass beads, as above.
Protocol 1. Samples are collected by filtration (as above) or collected by centrifugation (cultures only) in 2.0 ml bead beater tubes (Biospecs). To this, 1 ml L6 lysis buffer is added together with eight scoops (approx 250 ~1
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packed volume) of muffled glass beads. The samples can be frozen in liquid nitrogen and stored at -80°C until analysis. 2. Samples are thawed and immediately disrupted by bead beater treatment for 2 min. The samples are centrifuged for 5 rain at 10 000 rpm at room temperature in a microcentrifuge. 3. The supernatant is collected, 70 btl of the silica solution is added, and the mixture vortexed. The pellet is re-extracted with an additional 1.0 ml of L6 buffer and 2 min of bead beating, followed by centrifugation. To the supernatant, 70 btl of the silica solution is added. 4. The silica-treated extracts are vortexed every 2rain for a total of 10 min. 5. The extracts are centrifuged for 15 s at 14 000 r p m in a microcentrifuge. This is most easily accomplished by holding the 'Pulse' switch of the microcentrifuge for a total of 25 s. The supernatants are discarded, 1 ml of L2 wash buffer is added, and the sample is vortexed. 7. Steps 5 and 6 are repeated. 8. The silica is then taken up in 1.0 ml of ethanol, and vortexed. This mixture is microcentrifuged as above. 9. The supernatant is carefully discarded and 1.0 ml of acetone is added. The mixture is vortexed and microcentrifuged as above. 10. The supernatant is discarded and the silica dried in a heating block set to 56°C for 10 min with the lid off the tube. 11. The RNA is eluted from the silica by addition of 100 btl of DEPCtreated, I mM EDTA (pH 7.0). Alternatively, DEPC-treated DI m a y be used. This mixture is heated at 56°C for 10 rain with vortexing after 5 min. 12. The mixture is microcentrifuged for 2 rain at 10 000 rpm. 80 btl of the supernatant is transferred to a new tube, and microcentrifuged for 2 min at 10 000 rpm. 13. 70 ~1 of the supernatant is collected and used immediately or stored at -80°C. .
RNeasy R Mini protocol Qiagen has developed a series of kits for nucleic acid extraction from a variety of tissue or cell types. We have used the RNeasy R kit for RNA extractions of environmental and culture samples, although we have not exhaustively determined the relative efficiency of extraction nor the specificity of RNA extraction. We have found equivalent relative hybridization intensities when compared to the GIPS extraction m e t h o d using natural populations. What appears below is our adaptation of the method.
Equipment and supplies • •
•
Bead-beater, muffled glass beads, and sterile 2 ml vials as per the GIPS method above. RNeasyRMini Kit. ~-mercaptoethanol, 100% ethanol.
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Procedure 1. Samples are either collected by filtration onto sterile 25 m m 0.45 [am Durapore (Millipore) filters or the samples are collected by centrifugation (cell cultures). 2. While filtration or centrifugation is occurring, add 0.01 v o l u m e of [3mercaptoethanol to the RLT buffer (from RNeasy ~<Mini Kit): 600 ~tl is required per extraction. 3. Roll the filter and add it to a 2.0ml sterile bead-beater tube that contains 0.5 g of muffled glass beads and 600 ~tl of RLT buffer + [3mercaptoethanol. If using a cell culture, re-suspend the pellet in 600 ~tl of RLT buffer + [3-mercaptoethanol and add to a 2.0 ml sterile beadbeater tube containing 0.5 g of glass beads. Label the tube and immediately freeze in liquid nitrogen. The samples can n o w be stored at -80°C until extraction. 4. Thaw the pellet or filter tube quickly by immersing in w a r m (40-50°C) water for 5 rain. Disrupt by bead-beating for 2 min and then chill on ice for 5 min. Repeat for a total of three times (three rounds of beadbeating and icing). 5. Centrifuge in a microcentrifuge at 14 000 rpm at 4°C for 5 min. 6. Transfer lysate (N450 ~tl) to a new tube and centrifuge for 2 min at 14 000 rpm at 4°C. Transfer supernatant to a new tube. 7. Add 0.7 v o l u m e (-315 gl) of 100% ethanol and mix by trituration. 8. Place sample including any precipitate which m a y have formed on an RNeasy mini spin colunm sitting in a 2 ml collection tube (supplied in kit). Centrifuge for 15 s at 10 000 rpm. If the v o l u m e >700 ~tl, two separate centrifugations can be performed. The collection tube can be reused but discard the liquid collected after each spin by emptying the collection tube. 9. Pipette 700 ~tl of buffer RW1 (RNeasy kit) onto the RNeasy column used in step 8. Centrifuge at 10 000 r p m for 15 s to wash. Discard the collection tube containing the wash. 10. Attach a new 2 ml collection tube (supplied in kit). Pipette 500 gl of RPE buffer (in kit) onto the column and centrifuge for 15 s at 10 000 rpm. Be certain that ethanol has been a d d e d to the RPE buffer, and see notes in the booklet accompanying the manufacturer's kit. 11. Pipette an additional 500 ~tl of RPE buffer onto the RNeasy column and centrifuge for 2 min at m a x i m u m speed (14 000 rpm) to dry the RNeasy membrane. It is important to dry the RNeasy m e m b r a n e since residual ethanol may interfere with subsequent steps. Carefully remove the 2 ml collection tube being certain not to contact the column with ethanol. 12. Transfer the column to a new 1.5 ml collection tube (supplied with kit). Pipette 51 gl of RNase-free water (in kit) directly onto the RNeasy membrane. Let it stand for 1 rain, followed by centrifugation for 3 rain at 10 000 rpm. The collected material (usually 36 ~l) is the RNA fraction, which can be stored frozen at -80°C.
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DOTTING, PROBING,AND QUANTITATING mRNA LEVELS There are several concerns when interpreting mRNA levels of particular target genes in the mixed microbial assemblages that occur in aquatic environments. Most important are issues of specificity and fidelity. It is important to have some understanding of how specific the probes are that are to be employed. Do they target the organism or groups of organisms desired? In terms of fidelity, are there organisms that are of interest that may not hybridize as strongly to the probe as others? Each set of probes should be evaluated with known organisms in culture to answer these questions. We have taken the approach of using full length or near full length gent probes to detect transcription. For work on the RuBisCO large subunit gene, we have used different genes to detect the Form IB clade and the Form ID clade of this enzyme. For the Form IB, we have used the rbcL gene from Synechococcus PCC6301 and for Form ID, that of Cylindrotheca N1 (Pichard et al., 1997). These probes have been shown to hybridize to their respective algal groups (Pichard et al., 1997). To make such probes, the inserts were subcloned into vectors containing the Sp6 and T7 RNA polymerase promoters (we have used the Promega vectors pGEM3Z and 4Z but many others are currently available). To detect mRNA signals we employed antisensense transcript probes labeled with ~S-UTP (New England Nuclear) and to ensure that extracts were not contaminated with target DNA, samples were split and probed with the sense probe (Pichard et al., 1993). Because we never obtained hybridization to RNA extracts with sense probes, we deleted this control in our studies starting in 1997. However, we do recommend checking extracts for DNA contamination by nuclease digestion, particularly when employing the Boom and RNeasy protocols as these have either shown a lot of DNA liberation or have not been tested (see below). For nuclease digestion, we divide the final RNA solution into three aliquots, leaving one untreated, and digesting one each with DNase (RNase-free, such as RQI DNase, Promega) and RNase, which has been heat-treated to remove DNase activity. Magnesium chloride is added to facilitate nuclease digestion. If the RNeasy protocol is used, we divide the 36 ~1 eluate into three 12 pl aliquots and treat as below:
Treatment
Undigested DNAse digested RNAse digested
RNAsin
2 M MgCI 2
RQI DNase
RNAse
Total volume
(,I)
(~l)
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While keeping the undigested sample on ice, the digested samples are incubated for 2 h at room temperature. 80 ~1 of DI water from the kit is added to the undigested and DNAse-digested samples. These samples are 404
immediately dotted onto charged nylon filters using a dot-blot apparatus. The filter is then removed from the dot-blot apparatus and dried thoroughly. The RNAse digested sample is then dotted by hand, 1 pl at a time. This prevents contamination of the dot or slot blotter with RNAse. The filter is then dried, cross-linked using a UV-cross-linker, and stored at -20°C in a zip-lock bag until probed. Standard curves. We have found it convenient to make mRNA standard curves of the types of target genes to be probed by in vitro transcription using Riboprobe constructs and the sense promoter. The RNA is then quantitated using a RiboGreen RNA quantitation kit (Molecular Probes) and dotted in a dilution series, starting with 5 ng, and ending with 0.1 pg. Standard curve blots are made en masse, and stored at -20°C until needed. Quantitating results. We have had success quantitating unknowns with standard curves using a BioRad Model 363 Molecular Imager. This instrument works by use of screens that are exposed to the radioactivity, and then are scanned by an infrared laser to detect where the screen was 'exposed'. Exposure times are of the order of minutes to overnight for very low signals. The detection limit is usually ~0.5 pg.
A P P L I C A T I O N S A N D P E R F O R M A N C E OF T H E ASSAY We have not as yet performed systematic comparisons of all three extraction methods. We have performed a few side-by-side comparisons of the GIPS method with both the Boom and RNeasy methods, and we have most experience with the GIPS method. Extraction of natural populations of phytoplankton from Ft. Desoto, St Petersburg, FL and probed with the Form ID probe indicated that the Boom protocol resulted in hybridization that was at least as great as that with the GIPS reagent (unpublished data). A series of experiments were performed with E. coil F+amp (obtained from Mark Sobsey, University of North Carolina, Chapel Hill) in which exponentially-growing cells were extracted by both the GIPS and Boom methods and both RNA and DNA were quantitated by fluorometric assays. At high cell abundance (20 ml culture N1-2 x 10" cells extracted) the GIPS method yielded greater levels of RNA, whereas at lower cell abundance (5 ml cells extracted) the Boom method yielded greater RNA. It may be that the silica was saturated with nucleic acid at the high cell densities. The RNeasy has been compared once with the GIPS method for natural populations and equivalent hybridization was found.
FUTURE DIRECTIONS 1 forsee the need to adapt current gene expression technology, which is still relatively cumbrous, to rapid, near real-time, small-scale mRNA detection. A drawback of current technology is the lack of sensitivity of direct probing which requires large volumes of samples to be filtered. One 405
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avenue of investigation we are pursuing is the amplification of message to increase sensitivity. One such approach is the use of Nucleic Acid Based Sequence amplification (NASBA; Davey and Malek, 1989; Van Gemen et al., 1994; Paul et al., 1999a). This method has been applied to a series of RNA targets. However, this procedure is perhaps best suited to detection of low levels of mRNA targets, but m a y not be applicable to quantitation of those targets. An alternative to that approach is what has been termed real-time PCR, using instrumentation developed by PE Applied Biosystems. Using this technology, RT-PCR is performed and the cycle at which exponential amplification occurs is indicative of initial target concentration. Therefore, it m a y be possible to rapidly extract RNA from a small volume (requiring filtration of only a few milliliters) followed by rapid, specific quantitation of mRNA targets. A second area of tremendous future potential is the application of DNA microarrays (gene chips) to problems concerned with the detection of environmental gene expression (Sterling, 1999; Marshall and Hodgson, 1998). Currently, microarrays are used to assay gene expression in one organism (say yeast or ovarian cancer cells) by having thousands of gene probes immobilized on a surface from that one organism. An RNA extract is made followed by an RT fluorescent labeling step, and the extract hybridized to the array. The identity of genes being expressed is determined by their position in the array. Arrays of environmentally important genes could be constructed, and the types of organisms and genes being expressed could potentially be determined by judicious probe selection. Such technology could be adapted eventually to be installed in remote moorings to provide near real-time gene expression analysis in situ.
References Bartlett, D. H. and Welch, T. J. (1995). ompH gene expression is regulated by multiple environmental cues in addition to high pressure in the deep-sea bacterium Photobacterium species strain SS9. J. Bacteriol. 177, 1008-1016. Boom, R., Sol, S. J., Salimans, M, M., Jansen, C. L., Wertheim-van Dillen, P. M. and van der Noordaa, J. (1990). Rapid and simple method for purification of nucleic acids. J. Clin. Microbiol. 28, 495-503. Chomczynski, P. and Sacchi, N. (1987). Single-step method of RNA isolation by acid guanidinium isothiocyanate-phenol-chloroform extraction. Anal. Biochent. 162, 156-159. Chow, T.-J. and Tabita, E R. (1994). Reciprocal light-dark transcriptional control of nif and rbc expression and light-dependent posttranslationaI control of nitrogenase activity in $ynechococcus sp. strain RF-1. ]. Bacteriol. 176, 6281-6285. Davey, C. and Malek, L. T. (1989). Nucleic acid amplification process. European Patent No. EP0329822. Hamamoto, T. I., Takata, N., Kudo, T. and Horidoshi, K. (1995). Characteristic presence of polyunsaturated fatty acids in marine psychrophilic vibrios. FEMS Microbiol. Lett. 129, 51-56. MacGregor, B. J. (1999). Molecular approaches to the study of aquatic microbial communities. Curt. Opin. Biotech. 10, 220-224. Marshall, A. and Hodgson, J. (1998). DNA chips: an array of possibilities. Nat. Biotech. 16, 27 31.
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Mate, Z., Sass, L., Szekeres, M., Vass, 1. and Nagy, F. (1998). UV-B-induced differential transcription of psbA genes encoding the D1 Protein of Photosvstem 11 in the cyanobacterium Sym'chocystis 6803.1. Biol. Chem. 273, 17439-17444. Paul, J. H., Ewert, M., Wawrik, B. and Stokes, R. (1999a). Novel RNA technology for microbial detection in aquatic environments. American Society for Limnology and Oceanography Ocean Sciences Meeting, Santa Fe, New Mexico. Abst #78, session SS41. Paul, J. H., Pichard, S. L., Kang, J. B., Watson, G. M. F., and Tabita, F. R. (1999b). Evidence for a clade-specific temporal and spatial separation in ribulose bisphosphate carboxylase gene expression in phytoplankton populations off Cape Hatteras and Bermuda. Limmd. Oceam~g. 44, 12-23. Pichard, S. L. and Paul, ]. H. (1991). Detection of gene expression in genetically engineered microorganisms and natural phytoplankton populations in the marine enviromnent by mRN A analysis. Appl. Envirott. Microbiol. 57, 1721-1727. Pichard, S. L., Frischer, M. E. and Paul, J. H. (1993). Ribulose bisphosphate carboxylase gene expression in subtropical marine phytoplankton populations. Mar. Ecol. Prog. Set'. 101, 55-65. Pichard, S. L., Campbell, L., Carder, K., Kang, J. B., Patch, J., Tabita, F. R. and Paul, J. H. (1997). Analysis of ribulose bisphosphate carboxylase gene expression in natural phytoplankton communities by group-specific gene probing. Mar. Ecol. Pr0~. Ser. 149, 239-253. Robinson, K. A., Robb, E T. and Schreier, H. J. (1994). Isolation of maltoseregulated genes from the hypertherlnophilic archaeum, Pyrococcus furiosus by subtractive hybridization. Gem' 148, 137-141. Ro> S. K. and Nakamoto, H. (1998). Cloning, characterization, and transcriptional analysis of a gene encoding an alpha-crystalline-related, small heat shock protein from the thermophilic cyanobacterium Syncchococcus vulcauus. I. Bocteriol. 180, 3997-4001. Sadeghi, A. (1998). Gene expression in two cyanobacteria, freshwater Syttechococcus PCC7942 and oceanic Sym'chococcus sp. WH7803, in response to ammonium, nitrate or iron. Dissert. Abstr. hH. 59, 5. Sambrook, ]., Fritsch, E. E and Maniatis, T. (1989). Molecular Clonitt,~: A Laboratory Mamtal, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor. Sterling, J. (1999). French groups exploit novel use of GeneChip. Gcw. Ensqm'et" News 19, 1-10. Techkarnjanaruk, S. and Goodman, A. E. (1999). Multiple genes involved in chitin degradation from the marine bacterium Psemtoalteromonas sp. Strain $91. Microbiol. 145, 925 934. van Gemen, B., van Beuningen, R., Nabbe, A., van Strijp, D., Jurriaans, S., Lens, P. and Kievits, T. (1994). A one-tube quantitative HIV-1 RNA NASBA nucleic acid amplification assay using electrochemilummescent (ECL) labeled probes. I. Vir. Methods 49, 157-168. Wilson, M. S., Bakermans, C. and Madsen, E. L. (1999). In situ, real-time catablic gene expression: extraction and characterization of naphthalene dioxygenase mRNA transcripts from groundwater. Appl. Envirom Microbiol. 65, 80 87. Wyman, J., Zehr, J. P., and Capone, D. G. (1996). Temporal variability in nitrogenase gene expression in natural populations of the marine cyanobacterium Trichodesmium thiebautii. Appl. Ettviron. Microbiol. 62, 1073-1075. Wyman, M. (1999). Diel rhythms in ribulose-l,5-bisphosphate carboxylase/oxygenase and glutamine synthetase gene expression in a natural population of marine picplanktonic cyanobacteria (Syttechococcus spp.). Appl. Ei1virolt. Microbiol. 65, 3651-3659.
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List of suppliers Bel-Art Products 6 industrial Road Pequannock, NJ 07440-1992, USA Tel: 1-800-423-5278 Fax: 1-800-545-9796 Website:www.belart.com Portable fume hoods
Molecular Probes, Inc. 4849 Pitchford Avenue POBox 22010 Eugene, OR 97402-0414, USA Tel: 1-800-438-2209 Fax: 1-800-438-0338 Website: www.probes.com Nucleic acid fluorochromes
Bio-Rad Laboratories Life Sciences Group 2000 Alfred Nobel Drive Hercules, CA 94547, USA Teh 1-800-424-6723 Fax: 1-800-879-2289 Website: www.bio-rad.com
Molecular Research Center 5645 Mon tgmneray Road Cincinnati, OH 45212, USA Tel: 1-888-841-0900 Fax: 1-513-841-0080 Website: www.mrcgene.com RNA extraction kit (Tri-Reagent)
Molecular Imaging instrumentation
Organon Teknika BV Bosund 15, PO Box 84 5280 AB Boxtel The Netherlands Tel: 31-411-654-931 Fax: 31-411-654-311 Website: www.nuclisens.com
Biospec Products PO Box 722 Bartlesville, OK 74005, USA Teh 1-918-336-3363 Fax: 1-918-336-3363 Website: www.biospec.com Bead-beating instrumentation and supplies Fisher Scientific Projects Division 50 Fadem Road Springfield, NJ 07081-3193, USA Tel: 1-800-766-7000 Fax: 1-800-926-1166 Website: www.fishersci.com Scientific and biotechnology supplies Millipore Corporation 80 Ashby Road Bedford, MA 01730, USA Tel: 1-800-645-5476 Fax: 1-781-275-5550 Website: www.millipore.com
RNA extraction kits Qiagen, Inc. 28159 Avenue Stanford Valencia, CA 91355-1106, USA Teh 1-800-426-8157 Fax: 1-800-718-2056 Website: www.qiagen.com RNeasy kits Sigma Chemical Co. 3050 Spruce Street St. Louis, MO 63103, USA Tel: 1-800-325-3010 Fax: 1-800-325-5052 Website: www.sigma-aldrich.com DEPC and other chemicals
Filtration supplies
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21 In situ PCR/RT-PCR Coupled with in situ Hybridization for Detection of Functional Gene and Gene Expression in Prokaryotic Cells ¢O
Feng Chen' and R o b e r t E Hodson 2 ~Center of Marine Biotechnology,University of Maryland BiotechnologyInstitute,Baltimore,MD 21202, USA;2Department of Marine Sciences,University of Georgia, Athens,Georgia 30602-2206, USA
CONTENTS
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Introduction Principles of PI-PCR Chemicals, enzymes and supplies In situ PCR for detecting genes In situ RT-PCRfor detecting mRNA targets In situ hybridization Other technical comments Conclusion
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INTRODUCTION Monitoring microbial genetic diversity and function at the single-cell level is critical for understanding the ecological role of microorganisms in aquatic environments. Fluorescent in situ hybridization (FISH) methods with phylogenetic probes have been widely used to identify specific prokaryotic cells in various natural communities (DeLong et al., 1989; Amann et al., 1990, 1995; Ramsing et al., 1996). A major limitation of the FISH technique is that natural bacterioplankton may contain insufficient cellular rRNA to yield a detectable fluorescent signal. Due to low copy number of messenger RNA (mRNA), it is even more difficult to detect the expression of functional genes inside prokaryotic cells using FISH. FISH is a straightforward and established method, which is mainly used for taxonomic or phylogenetic identification of bacterial cells within microbial communities. Like 16S rDNA, the DNA sequence information of a functional gene can be used to describe phylogenetic relationships between microbes
METHODS IN MICROBIOLOGY, VOLUME 31} 1SBN (/ 12-521530-4
Copyright © 20(}1 Academic Press Ltd All rights of reproduction in any form reserved
sharing a common phenotype. The presence of a given functional gene is generally associated with a particular bacterial metabolic capability. Polymerase Chain Reaction (PCR) technology has been extensively used to amplify and detect the presence of several functional genes in aquatic microbial communities, such as the genes responsible for nitrification (Sinigalliano et al., 1995; Rotthauwe et al., 1997), denitrification (Ward et al., 1993; Scala and Kerkhof, 1998, 1999), nitrogen fixation (Kirshtein eta/., 1991; Zehr et al., 1995, 1998), and carbon fixation (Paul and Pichard, 1998). Many microbial genes from natural aquatic environments have been amplified and sequenced. Consequently, new PCR primers can be designed to amplify and detect specific functional genes for more specific microbes in natural habitats. For many years, microbial ecologists have been seeking to develop techniques to better understand the function and diversity of microbial communities in natural marine environments, where we presume less than 1% of microbes to be cultivable. Given that a functional gene represents the potential capability for a specific microbial activity, it is generally assumed that gene expression or messenger RNA (mRNA) production induced by environmental changes can be used as an indicator for the actual microbial activity associated with this gene. mRNA extracted from microbial communities have been detected by reverse transcription (RT)PCR amplification (Pichard et al., 1997; Miskin et al., 1999) or by hybridization to specific radioactively-labeled probes (Pichard and Paul, 1993). RT-PCR is a very sensitive method for detecting minute quantities of mRNA. PCR or RT-PCR performed against extracted DNA or RNA from natural communities is often called in vitro PCR or RT-PCR. In vitro PCR/RT-PCR performed on bulk extraction of nucleic acids is useful for detecting the presence of genes (or gene fragments) that are associated with a specific function of a microbial commmlity. However, this approach cannot provide information on the abundance and distribution of microbial cells with a particular metabolic activity. Nearly ten years ago, a technique called in situ PCR was successfully used to amplify and detect viral DNA inside eukaryotic cells (Bagasra, 1990; Haase et al., 1990). Since then, in situ PCR methods have been widely used in medical research and undergone rapid development and modification (Bagasra et al., 1995; Gu, 1995; Gosden, 1997). hz situ PCR has been used to specifically amplify and detect single copy or low copy nucleic acid sequences in single cells and tissue sections (Long and Komminoth, 1995). More recently, Hodson et al. (1995) developed a Prokaryotic in situ PCR (PI-PCR) method to visualize the microscale distribution of specific genes and gene products in individual bacterial cells in microbial communities. In the past several years, various P1-PCR protocols have been developed to investigate functional gene occurrence and gene expression inside bacterial cells (Porter et al., 1995; Chen et al., 1997, 1998, 1999; Tolker-Nielsen et al., 1997, 1998; Jacobs et al., 1997; Tani et al., 1998; Kurokawa et al., 1999; Holmstrom et al., 1999). PI-PCR protocols have been used to detect naphthalene degradation (nahA) and toluene degradation (todC1) genes in Pseudomonas cells and in model marine bacterial communities (Hodson et al., 1995; Chen et al., 1998, 1999). P1-PCR and PI-RT-PCR 410
have been coupled with flow cytometry to quickly enumerate bacterial cells with a specific functional gene (Porter et al., 1995) and gene expression (Chen et al., 2000). PI-RT-PCR with labeled primers has demonstrated the potential to detect poorly expressed mRNA in individual bacterial cells (Tolker-Nielsen et al., 1997), and this approach has been used to detect the expression of several functional genes in individual Salmonella typhimurium cells (Tolker-Nielsen et al., 1997; Holmstrom et al., 1999). Direct PI-PCR with a novel dye, HNPP/Fast Red TR, was used to detect 16S ribosomal DNA of E. coli and thus identify cells specifically at the single-cell level in a bacterial mixture (Tani et al., 1998). A similar method was able to quantify those bacteria carrying a specific functional gene in river water (Kurokawa et al., 1999). These studies indicate that PI-PCR methods can provide microbial ecologists with new insights into the genetic diversity and activity of microbes at the single-cell level. Although there are a variety of PI-PCR protocols available, PI-PCR coupled with FISH has been found to provide a higher specificity than other protocols (Chen et al., 1999). For PI-PCR with direct incorporation of labeled nucleotides, false positives resulting from non-specific incorporation of labeled nucleotides can be significant. Several factors may contribute to a false positive signal: (1) incorporation of labeled nucleotides into cellular DNA by the repair activity of DNA polymerase; (2) mispriming, in which added primers bind to non-target sequence regions; (3) endogenous priming, in which endogenous DNA or RNA fragments act as primers for PCR amplification; (4) unstable binding between fluorochrome and dUTP during heating and cooling cycles of PCR which releases fluorochrome resulting in a general staining of all cells; or (5) binding of fluorecently-labeled dUTP to the cellular components inside cells due to high temperatures reached during PCR. Some of these problems cannot be eliminated even if a second nested i, situ PCR is used. The combination of PI-PCR and FISH greatly reduces the false positive signal because the initial i, situ PCR/RT-PCR is performed with nonlabeled nucleotides. Additionally, the stringency of FISH can be critically optimized by varying hybridization conditions (e.g. temperature, formamide and salt) to achieve more specific detection with low background. Recent studies of eukaryotic i, situ PCR suggested that i, situ PCR coupled with FISH provides much higher specificity than direct in situ PCR (Nuovo, 1992; Zehbe et al., 1994; Long and Komminoth, 1995; Bagasra et al., 1995). Most recently, PI-PCR coupled with FISH was exploited to monitor expression of the todC1 gene in Pseudomo,as putida grown in seawater exposed to toluene (Chen et al., 1999). In this chapter, we will provide detailed protocols for performing PIPCR on microscope slides, including prokaryotic iJl situ PCR, in situ RTPCR and in situ hybridization.
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PRINCIPLES
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PI-PCR differs from eukaryotic ill situ PCR in many aspects: (1) prokaryotic cells are usually too small (less than a few micrometers) for iJ~ situ 411
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PCR/RT-PCR to localize genes and their transcripts in intact ceils under epifluorescence microscopy; (2) prokaryotic mRNAs usually have much shorter half-lives (< 10 min), therefore, rapid fixation and optimal preservation are necessary to detect mRNA transcripts using PbRT-PCR; (3) cell wall permeabilization by lysozyme is critical for successful PI-PCR amplification. Unlike in vitro PCR/RT-PCR in which extracted DNA or RNA is used as a template for amplification, PI-PCR is performed using DNA or RNA inside bacterial cells where the cell membrane acts as a container for amplified products. The nature of the cellular material containing the target sequences to be amplified is an important variable for PI-PCR. Successful in situ PCR amplification depends on extensive optimization
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procedures to determine the conditions which will permeabilize the bacterial cell membranes to allow entry of reagents for amplification and detection, yet retard the diffusion of PCR product out of the cells. If the amplification target is DNA, cellular RNA should be digested with DNase-free RNase (Figure 21.1). However, if the amplification target is RNA (rRNA or mRNA), permeabilized cells are treated with RNase-free DNase (Figure 21.2). Performing in situ RT and PCR reactions in one mixture using the specific enzymes, such as rTth DNA polymerase (PE Applied Biosystems) or Titan TM One Tube RT-PCR system (Roche Molecular Biochemicals) has greatly simplified in situ RT-PCR procedures, Currently, there are two different methods for detecting amplified PCR products inside cells, direct or indirect in situ PCR. For direct PI-PCR,
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that optimal procedures can be established and adapted to suit specific needs.
CHEMICALS, ENZYMES A N D SUPPLIES • Gelatin solution: 0. I% gelatin (Sigma Chemical Co.), 0.0 I% chromium potassium sulfate (Sigma Chemical Co.) • Ix phosphate buffered saline (PBS, Sigma Chemical Co., Cat. No. P 8033): 120 mM NaCI, 2.7 mM KCI, I 0 mM phosphate buffer salt, pH 7.6 • 8% paraformaldehyde (Sigma Chemical Co.) in PBS • Ethanol series: 50, 80 and 98% (v/v) ethanol • Lysozyme stock (20 mg ml '):freshly prepared. Add 10 mg of lysozyme (Roche Molecular Biochemicals, Cat. No. 107255) to 500lul of lysozyme buffer (I 00 mM Tris, 50 mM EDTA, pH 8.0). • DNase-free RNase I: 500 lug ml' (Roche Molecular Biochemicals, Cat. No. II 19915) • RNase-free DNase I: 10U lul ~ (Roche Molecular Biochemicals, Cat. No. 776785) • RNase-free DNase solution: 40 mM Tris-HCI, pH 7.4, 6 mM MgClv 2 mM CaCI2 • Proteinase K stock: freshly prepared. Add 40 lug of proteinase K (Roche Molecular Biochemicals, Cat. No. 161519 or 745723) to I ml of PBS. • 0.5x SSC: 75 mM NaCI, 7.5 mM trisodium citrate, pH 7.0 • DEPC treated dH20: add I ml of DEPC to 1000 ml of dH20 (prepare in hood), incubate at 37°C for 2 h, then autoclave the treated dH20. • RNase inhibitor:40 U lul' (Roche Molecular Biochemicals, Cat. No. 799017) • Hybridization solution: 360 lul 5M NaCI 40 lul I M Tris-HCI pH 8.0 x lul formamide (according to the Tm of probe) Adjust volume to 2 ml with distilled water I lul 10% SDS • Washing buffer: I ml I MTris-HCI pH 8.0 (I M) S00 lul 500 mM EDTA y lul % 5 M NaCI (according to the amount of formamide used) Adjust volume to 50 ml with distilled water 50 lul 10% SDS • Counter-staining solutions: DAPI (Sigma Chemical Co., Cat. No. D-9542) and SYBR Green I (Molecular Probes, Inc., Cat. No. S-7563) • Expand TM High Fidelity PCR System (Roche Molecular Biochemicals, Cat. No. 17364 I) • Titan TM One Tube RT-PCR system (AMV and Expand TM High Fidelity PCR System, Roche Molecular Biochemicals, Cat. No. 1888382 or 1855476) • PTC-200Thermal Cycler with the tower unit for slides (MJ Research) • Moist chambers (to place slides for in situ hybridization) • Incubation oven (for in situ hybridization) • Microscopic slides (Gold Seal Products, Cat. No. 3049) • Coplin jars • Frame-seal chamber with polyester covers (MJ Research, Inc., Cat. No. SLF0201,9 × 9 ram)
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• Microcentrifuge (Eppendorf Model 5414C) • Epifluorescent microscope: with excitation wavelengths at 350, 496, and 596 nm. • Digital cooled CCD camera (optional) • Digital image analysis system (optional)
Coating microscope slides with gelatin 1. Gently wipe microscope slide surface with 100% ethanol and allow to air dry. 2. Warm the gelatin solution in oven or incubator (50°C) and transfer the solution into a Coplin jar or a glass staining dish. 3. Dip the clean slide into the w a r m gelatin solution for 1 min. 4. Pull the slide out of Coplin jar and allow to air-dry in a vertical position for a few hours or overnight. 5. Store gelatin-coated slides in a sealed container at room temperature, and use them within two weeks.
Preparation of 8% paraformaldehyde in PBS (40 ml) 1. Heat 25 ml of distilled water in a flask to nearly boiling (about 90°C) on a heating/stirring plate. 2. Drop a magnetic stir bar into the flask and start stirring 3. A d d 3.2 g of p a r a f o r m a l d e h y d e into the flask. 4. A d d one drop at a time of 1 N N a O H solution into the flask until the solution becomes clear. 5. Add 13 ml of 3× PBS into the flask and adjust p H to 7.6. 6. Filter the solution through a 0.2 lain filter, and store the paraformaldehyde solution at 4°C for no more than one month.
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Sample fixation and permeabilization 1. Add one v o l u m e of 8% p a r a f o r m a l d e h y d e to one v o l u m e of sample, and fix sample in ice for i h. 2. Pellet fixed cells by centrifugation (6000rpm, 3 min), and remove supernatant. 3. Wash cells twice with PBS using centrifugation (6000 rpm, 3 rain). 4. Re-suspend cells in PBS by gentle pipetting, add equal v o l u m e of 100% ethanol and mix. Adjust the re-suspended cells to a final concentration of 1if' cells mV. At this stage, the cells can be stored at -20°C for one year if used for PI-PCR. 5. Place a frame-seal chamber (two per slide) onto a clean gelatin-coated glass slide, and pipette 5 lJl of fixed cells (about 10: total cells) into the frame (9 m m × 9 mm). Dry the slide in the oven with temperature preset at 50°C. 416
6. Dilute lysozyme stock (20 mg ml ') to a working solution (2 mg ml ') using lysozyme buffer, and add 50 pl of lysozyme working solution onto each sample spot on the slide. 7. Incubate the slide for 15 min at room temperature. Rinse the slide with PBS once. 8. Add 50 pl of DNase-free RNase I (5 lag ml [, final concentration) onto each sample spot on the slide and incubate for 15 rain at room temperature. Rinse the slide with PBS once. 9. Cells on the slide are dehydrated in 50, 80 and 98% ethanol, sequentially, 1 min each. The slide is dried in a 50°C oven for 2 min, then add the PI-PCR cocktail to the sample spot.
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2. Add 25 lal of PI-PCR reaction mix into each sample on the slide, and carefully place a polyester cover slip over the frame. 3. Place slides into the tower block on the PTC-200 Thermal Cycler (MJ Research, inc.) and start PCR cycling. PCR amplification starts with an initial 3 min denaturation at 94°C, followed by 30 thermal cycles (denaturation at 94°C for 30 s, annealing at the primer-specific temperature for 30 s, and extending at 68°C for 30 s). 4. After PCR, remove the frame-seal assembly by holding d o w n the slide and pulling back the tab of the cover slip along the same plane as the slide. 5. Wash the slide twice with 0.Sx SSC buffer at room temperature, 5 min per wash. 6. Cells on the slide are dehydrated again in 50, 80 and 98% ethanol, sequentially, 1 rain each. Air-dry the slide. At this stage, the intracellular gene target amplified by PI-PCR can be detected by in situ hybridization (see the in situ hybridization protocol).
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•
IN SITU RT-PCR FOR D E T E C T I N G m R N A TARGETS
Sample fixation and permeabilization l. Fix cells in 4% p a r a f o r m a l d e h y d e for 1 h in ice and harvest fixed cells by centrifugation (6000 rpm, 3 rain). 2. Wash fixed cells twice with ice-chilled PBS using centrifugation (6000 rpm, 3 min). 3. Re-suspend the cell pellet in ice-chilled PBS by gentle pipetting, add equal v o l u m e of ice-chilled 100% ethanol. Adjust the fixed cells to a final concentration of 10" ml '. At this stage, fixed cells can be stored in E T O H / P B S (1:1) at-20°C for three months. 4. Place the frame-seal chamber (two per slide) onto a clean gelatincoated glass slide, and pipette 5 pl of fixed cells (about 10 ~ total cells) into the frame (9 m m × 9 mm). Dry the slide in a 50°C oven. 5. Add 50 I~l of lysozyme working solution (2 mg ml ~, final concentration) onto each sample spot on the slide and incubate for 15 min at room temperature. Rinse the slide with PBS once. 6. Add 50 t-fl of RNase-free DNase I (0.1 U lJl ', final concentration) onto each sample spot and incubate the slide for 2 h in a moist chamber at room temperature. Inactivate DNase activity by heating the sample at 95°C for 5 rain. Rinse the slide with PBS once. 7. Cells on the slide are then d e h y d r a t e d in 50, 80 and 98% ethanol, sequentially, 1 min each. Slide is dried in a 50°C oven for 2 min. It is n o w ready for the PI-PCR cocktail.
Performing PI-RT-PCR on slides 1. PI-RT-PCR mixture (100 pl total): DEPC-treated dH,O 5x reaction buffer with 7.5 mM Mg >* 10x d N T P mix (2.5 raM) Upstream primer (100 lJM) Downstream primer (100 pM) MgCI: (25 raM) ~ Titan TM One Tube RT-PCR enzymes ~ RNase inhibitor (40 U pl ') DTT (100 mM) ~
55 lal 20 I~1 10 lJl 1 pl 1 pl 5 pl 2pl I ~21 5 V1
~These p r o d u c t s come with tilt, Titan ~~ One Tube RT-PCR system. A d d i t i o n a l a m o u n t of Mg ~ is a d d e d to increase the efficiency of PI-RT-I'CR. The final concentration of M g ' ill the PI-RT-PCR mixture is 2.75 m~l.
Add 25 1J1 of the PI-PCR reaction mix to each sample on the slide, and carefully place a polyester cover slip over the frame. 3. Place slides into the tower blocks on the PTC-200 Thermal Cycler (MJ Research, Inc.) and start RT-PCR cycling. Reverse transcription is performed at 60°C for 30 min. PCR amplification starts with an initial 2.
418
3 min denaturation at 94°C, and followed by 30 thermal cycles (denaturing at 94°C for 30 s, annealing at the primer-specific temperature for 30 s, and extension at 68°C for 30 s). 4. After RT-PCR, remove the frame-seal assembly by holding d o w n the slide and pulling back the tab of the cover slip along the same plane as the slide. 5. Wash the slide twice with 0.5x SSC buffer at room temperature, 5 min
per wash. 6. Dehydrate cells on the slide again in 50, 80 and 98% ethanol, sequentially, 1 rain each. Air-dry the slide. At this stage, the intracellular mRNA target amplified by PI-RT-PCR can be detected by in situ hybridization with a fluorescently labeled probe (see the in situ hybridization protocol). Notes: Prokaryotic mRNA is very sensitive to RNase activity and degrades quickly. Therefore, fresh cultures or samples should be fixed immediately with 4% paraformaldehyde at the point of sampling. We recommend that the sample pre-treatment be conducted at low temperature as much as possible to minimize the RNase activity. Water, reagent bottles, microcentrifuge tubes and glass slides should be pre-treated with DEPC. RNase inhibitor is added into RT-PCR mixture as a precaution against any RNase contamination from the other reagents.
IN SITU H Y B R I D I Z A T I O N 1. A d d 50 lal of hybridization solution containing 5 ng pl ~(final concentration) of a fluorescently labeled probe to each sample spot on the slide, and cover the spot with a cover slip. 2. Incubate slides at 90°C for 1-2 rain on the in situ PCR machine (to denature intracellular double-stranded DNA fragments amplified by PI-PCR or PI-RT-PCR), and then transfer to a h u m i d chamber preheated to the hybridization temperature required for the specific probe. 3. Incubate the slide in a h u m i d chamber at required temperature for 3 h or overnight. 4. After incubation, wash the slide with washing buffer for 10 min at a temperature slightly higher (2 to 3°C) than the probe hybridization temperature. 5. In order to visualize all bacterial cells (positive and negative) on the slide, cells should be counterstained with either 4',6'-diaminino-2phenylindole (DAPI) at a final concentration of 0.5 lag ml ' or a 1:1000 dilution of SYBR Green I nucleic acid stain. Cover tile sample spot with 1-2 drops of stain and incubate for 5 min at room temperature. 6. Rinse off the DAP1 or SYBR Green 1 from slide with distilled water. 7. Air-dry the slide and add a drop of emulsion oil onto the sample spot. Examine the slide u n d e r an epifluorescence cnicroscope (1000x).
419
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U'-
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U
Notes: Most of our primers and labeled probes were synthesized in custom synthesis laboratories (e.g. Sigma-Genosys or Life Technology). Various types of 5'-end labeling are available from these laboratories including biotin, digoxigenin and many other fluorescent dyes (e.g. fluorescein, Rhodamine and CY3). We recommend the labeled probe to be purified by HPLC or PAGE for in situ hybridization analysis. In most cases, in situ hybridizations are performed for three hours, however, overnight hybridization can be used for convenience. For probes with higher Tm, the formamide concentration can be increased in the hybridization buffer to maintain hybridization stringency at lower temperatures. In our laboratory, we use the program Oligo '~' 4.05 Primer Analysis Software (National Biosciences, Inc.) for determining the optimal hybridization conditions for a specific probe. Generally, we prefer to use a temperature around 50°C for in situ hybridization. The fluorescent dye, SYBR Green 1, is a bright fluorescent dye and stains both DNA and RNA. Therefore, it is more suitable for samples treated with DNase.
OTHERTECHNICAL
COMMENTS
Types of fixatives and length of fixation time are probably the most complicated issues for many in situ molecular detection methods. In a recent study we demonstrated that fixing bacterial cells in 4% paraformaldehyde for one hour on ice was sufficient to maintain cellular integrity throughout itl situ PCR amplification and hybridization. Moreover, in vitro RT-PCR was able to amplify a target sequence from extracted RNA of fixed cells which had been stored in ETOH/PBS (1:1) at -20°C for three months (Bachoon et al., in press). Prolonged fixation may cause cross-linking of nucleic acids with other cellular components which, in turn, can result in lower efficiency for PI-PCR amplification. High background fluorescence signal and false positives are the two most common problems encountered in direct PI-PCR. For direct PI-PCR, background signal is caused by the binding of labeled nucleotides onto the surface of the microscope slide between bacterial cells, while false positive results were attributed to the non-specific binding of labeled nucleotides onto cells. In contrast, indirect PI-PCR coupled with FISH detection significantly reduced the problems of high background and false positives (Chen et al., 1999). It should be noted that optimized conditions for in vitro PCR amplification may not always work for in situ PCR. In our laboratory, it was found that slightly increasing the concentrations of Mg ~', primers and enzymes tended to improve the performance of PIPCR. It is critical to ensure that the efficiency of PI-PCR is high and produces enough of the desired PCR product for FISH detection. It is possible to test the efficiency of intracellular amplification by analyzing the nucleic acids extracted from cells amplified by PI-PCR. One should carefully design and employ appropriate positive and negative controls for each specific PI-PCR experiment. Negative controls 420
of cells without DNA polymerase or without primers should always be included. For in situ amplification and hybridization, it is also important to use one slide as a control for non-specific binding of the probe, this can be done by hybridizing the PI-PCR cells with a probe for which there is no target DNA sequence in the cells. For PI-PCR, cells lacking the g e n t of interest, or cells treated with DNase, can be used as negative controls. For PI-RT-PCR, several negative controls can be chosen, such as cells treated with both DNase and RNase, cells not treated with reverse transcription, or cells in which the gene of interest is not expressed or induced. It is necessary to continuously validate the procedure and to confirm the efficiency of amplification because occasional technical pitfalls could cause misleading results. Although this chapter describes the protocols for performing PI-PCR on microscope slides only, bacterial cells in suspension are also suitable for PI-PCR. For instance, PI-PCR against suspended cells in a microcentrifuge tube has been coupled with flow cytometric analysis (Porter et al., 1995; Chert et al., 2000). However, the recovery efficiency of cells in suspension is usually lower than that for cells adhered to glass slides throughout the procedure. Our recent study found that approximately 80% of bacterial cells were recovered after PI-RT-PCR and FISH in bacterial cell suspension (Chert et al., 2000). We have used two different methods to concentrate bacteria from aquatic environments, centrifugation and filtration. The centrifugation method involves spinning fixed samples at 14000 rpm for 10 min, resuspending cells in PBS and preserving cells in ETOH/PBS (1:1) at -20°C. Bacterial densities in natural aquatic environments are usually in the range 105-10 ' ml '. Natural samples should be concentrated at least 1000-fold so that there will be sufficient bacterial cells to spread onto the microscope slide. Alternatively, bacterial cells in natural samples can be concentrated onto a membrane filter. Cells on the filter can be transferred onto a gelatin-coated slide, however, it is possible to perform PI-PCR directly against the cells on the filter. We have found that performing PIPCR with FISH directly against a bacterial smear on a slide usually yields less background noise than against bacterial cells on a membrane filter.
CONCLUSION The methods we provide here have been successfully used to detect the functional gene and gene expression inside bacterial cells of mixed culture, enriched microcosms and natural communities. However, it seems inevitable that new technologies like these will encounter new challenges when they are used to address ecological concerns in natural marine environments. I11 situ molecular detection of marine microbes will become more powerful when coupled with other advanced technologies such as digital image processing and flow cytometry. The ability to examine in situ microbial activities in terms of population distribution, 421
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genetic capability a n d function will p r o v i d e n e w insights on the i m p o r tant b i o g e o c h e m i c a l functions of m a r i n e microbial c o m m u n i t i e s .
Acknowledgments We a c k n o w l e d g e s u p p o r t f r o m the f o l l o w i n g US f u n d i n g agencies: Department of E n e r g y (DE-FG02-97ER62451), National Science F o u n d a t i o n (OCE-9730602 a n d OCE-0049098), N O A A / S e a Grants N a t i o n a l B i o t e c h n o l o g y P r o g r a m s (NA66RG0282), a n d the NSF L M E R p r o g r a m (DEB-9412089). C O M B c o n t r i b u t i o n No. 541
References Amann, R. I., Binder, B. J., Olson, R. J., Chisolm, S. W., Devereux, R. and Stahl, D. A. (1990). Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial population. Appl. Ellviron. Microbiol. 56, 1919-1925. Amann, R. I., Ludwig, W. and Schleifer, K.-H. (1995). Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59, 143-169. Bachoon, D., Chen, E and Hodson, E. E. fin press). RNA recovery and detection of mRNA by RT-PCR from preserved prokaryotic samples. FEMS Microbiol. Lett. Bagasra, O. (1990). Polymerase chain reaction in situ. Amplification March, 20-21. Bagasra, O., Seshamma, T., Pomerantz, R. J. and Hansen, J. (1995). lu situ PCR and hybridization to detect low-abundance nucleic acid targets. In: CurreI# Protocols iH Molecular Biology. Vol. 2. Sections 14.8.1-14.8.23. John Wiley and Sons, New York. Chen, E, Gonzalez, J. M., Dustman, W. A., Moran, M. A. and Hodson, R. F. (1997). In situ reverse transcription: an approach to characterize genetic diversity and activities of prokaryotes. Appl. EllviroH. MicJvbiol. 63, 4907-4913. Chen, E, Dustman, W. A., Moran, M. A. and Hodson, R. E. (1998). In situ PCR methodologies for visualization of microscale genetic and taxonomic diversities of prokaryotic communities. In: Moh'cular Microbial Ecology Mamml (A. D. L. Akkermans, J. D. van Elsas and E J. DeBruijn, Eds), pp. 1-17. Kluwer Acadamic Publishers, The Netherlands. Chen, E, Dustman, W. A. and Hodson, R. E. (1999). Detection of toluene dioxygenase gene and gene expression in Pseudomonas putida F1 in a toluene exposed seawater using in situ PCR and hybridization. Hydrobiologia 401, 131-138. Chen, E, Binder, B. J. and Hodson, R. E. (2000). Flow cytometric detection of specific gene expression in prokaryotic cells using in situ RT-PCR. FEMS Microbiol. Lett. 184, 291 295. DeLong, E. E, Wickham, G. S. and Pace, N. R. (1989). Phylogenetic stains: ribosomal RNA-based probes for the identification of single cells. Sciellce 243, 1360 1362. Gosden, J. R. (1997). PRINS and itl situ PCR protocols. In: Methods iu Molecular Biology, (J. M. Walker, Ed.) Vol. 7l. Humana Press, Totowa, NJ. Gu, J. (1995). In situ PCR - an overview-. In: In situ Polymerase Chain Reaction amt Related Technoh~gy (J. Gu, Ed.). pp. 1 2l. Eaton Publishing Co., Natick, MA. Hasse, A. T., Retzel, E. E and Staskus, K. A. (1990). Amplification and detection of lentiviraI DNA inside cells. Proc. Nat[. Acad. Sci. USA 7, I874-1878. Hodson, R. E., Dustman, W. A., Garg, R. P. and Moran, M. A. (1995). In situ PCR for
422
visualization of microscale distribution of specific genes and gene products in prokaryotic communities. Appl. Eilviron. Microbiol. 61, 4074-4082. Holmstrom, K., Tolker-Nielsen, T. and Molin, S. (1999). Physiological states of individual Salmouella typhimurium cells monitored by ilt situ reverse transcription-PCR. J. Bacteriol. 181, 1733-1738. Jacobs, D., Angles, M. L., Goodman, A. E. and Neilan, B. A. (1997). Improved methods for iH situ enzymatic amplification and detection of low copy number genes in bacteria. FEMS Microbiol. Lett., 152, 65-73. Kirshtein, J. D., Paerl, H. W. and Zehr, J. P. (1991). Amplification, cloning and sequencing of a nifH segment from aquatic microorganisms and natural communities. Appl. EnviroH. Microbiol. 57, 2645-2650. Kurokawa, K., Tani, K., Ogawa, M. and Nasu, M. (1999). Abundance and distribution of bacteria carrying sltlI gene in natural river water. Lett. Appl. Microbiol. 28, 405-410. Long, A. A. and Komminoth, P. (1995). hi situ PCR: General methodology and recent advances. In: In situ Polymerase ChaiH ReactioH amt Related Technology (J. Gu, Ed.), pp. 23-24. Eaton Publishing Co,, Natick, MA. Miskin, 1. P., Farrimond, P. and Head, I. M. (1999). Identification of novel bacterial lineages as active members of microbial populations in a freshwater sediment using a rapid RNA extraction procedure and RT-PCR. Microbiolo~y-UK 145, 1977-1987. Nuovo, G. J. (1992). PCR in situ Hybridizatioll: Protocols and Application,s. Raven Press, New York. Paul, J. H. and Pichard, S. L. (1998). Phytoplankton activity through the measuremerit of ribulose bisphosphate carboxylase gene expression (RuBisCO). In: Moh'czdar Approaches to the Study of the OceaH (K. E. Cooksey, Ed.), pp. 207-225. Chapman and Hall, London. Pichard, S. L. and Paul, J. H. (1993). Gene expression per gene dose, a specific measure of gene expression in aquatic microorganisms. Appl. E~r~,iro~l.Microbiol. 59, 451-457. Pichard, S. L., Campbell, L. and Paul, J. H. (1997). Diversity of the ribulose bisphosphate carboxylase/oxygenase form 1 gene (rbcL) in natural phytoplankton communities. Appl. EHvirom Microbiol. 63, 3600 3606. Porter, J., Pickup, R. and Edwards, C. (1995). Flow cytolnetric detection of specific genes in genetically modified bacteria using ill sitll polymerase chain reaction. FEM Microbiol. Lett. 134, 51 56. Ramsing, N. B., Fossing, H., Ferdehnan, T. G. Andersen, F. and Tbamdrup, B. (1996). Distribution of bacterial populations in a starified Fjord (Mariager Fjord, Denmark) quantified by ill sitll hybridization and related to chemical gradients in the water column. Appl. Elzviroll. Microbiol. 62, 139] 1404. Rotthauwe, J-H., Witzel K-L. and Liesack, W. (1997). The ammonia monooxygenase structural gene amoA as a functional marker: molecular fine scale analysis of natural ammonia-oxidizing populations. Appl. ElzviroJl. Microbiol. 63, 4704-4712. Scala, D. J. and Kerkhof, L. J. (1998). Nitrous oxide reductase O~osZ) gene-specific PCR primers for detection of denitrifiers and three HosZ genes from marine sediments. FEMS Microbiol. Lett. 162, 61-68. Scala, D. J. and Kerkhof, L. J. (1999). Diversitv of nitrous oxide reductase (nosZ) genes in continental shelf sediments. Appl. EnviroIT. Microbiol. 65, 1681-1687. Sinigalliano, C. D., Kuhn, D. N. and Jones, R. D. (1995). Amplification of the amoA gene from diverse species of ammonium-oxidizing bacteria and from an indigenous bacterial population from seawater. Appl. EJlviroH. Microbiol. 61, 2702-2706. Tani, K., Kurokawa, K. and Nasu, M. (1998). Development of a direct in situ PCR
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method for detection of specific bacteria in natural environments. Appl. Environ. Microbiol. 64, 1536-1540. Tolker-Nielson, T., Holmstrom, K. and Molin, S. (1997). Visualization of specific gene expression in individual Sahnonella typhimurium cells by in situ PCR. Appl. Environ. Microbiol. 63, 4196-4203. Tolker-Nielson, T., Holmstrom K., Boe, L. and Molin. S. (1998). Non-genetic population heterogeneity studied by in situ polymerase chain reaction. Mol. Microbiol. 27, 1099-1105. Ward, B. B., Cockcroft, A. R. and Kilpatrick, K. A. (1993). Antibody and DNA probes for detection of nitrite reductase in seawater. J. Gen. Microbiol. 139, 2285-2293. Zehbe, I., S~llstr6m, J. E, Hacker, G. W., Hauser-Kronberger, C., Rylander, E. and Wilander, E. (1994). Indirect and direct in situ PCR for the detection of human papillomavirus. An evaluation of two methods and a double staining technique. Cell Vision 163, 163-167. Zehr, J. P., Mellon, M. T., Braun, S. T., Steppe, T. and Paeral, H.W. (1995). Diversity of heterotrophic nitrogen fixation genes in marine cyanobacterial mat. Appl. Envilvn. Microbiol. 61, 2527-2532. Zehr, J. P., Mellon, M. T. and Zani, S. (1998). New nitrogen-fixing microorganisms detected in oligotrophic oceans by amplification of nitrogenase (nifH) genes. Appl. Environ. Micn~biol. 64, 3444-3450.
List of suppliers Gold SeaV Products
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424
22 Denaturing Gradient Gel Electrophoresis in Marine Microbial Ecology H e n d r i k Schiifer' and G e r a r d M u y z e r ~ 'Max-Planck-lnstitute for Marine Microbiology, Bremen, Germany; ~Department of Biological Oceanography, Netherlands Institute for Sea Research (NIOZ), Texel, The Netherlands
L9
CONTENTS
Introduction Practical aspects of PCR-DGGE Analysis of DGGE patterns Limitations of the PCR-DGGE approach Conclusions
• ~ ~
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INTRODUCTION During the past decade the approach to microbial community composition analysis has changed considerably. Classical techniques such as cultivation and microscopic identification are not sufficient to assess the diversity of bacteria in natural samples. On the one hand, lack of conspicuous morphology and small cell size do not allow microscopic identification of the majority of naturally occurring bacteria: on the other hand, media used for the cultivation of microbial strains are selective and hence give a biased view of the community composition. Furthermore, isolation of the vast majority of naturally occurring bacteria in pure culture is hindered by our lack of knowledge of the specific culture conditions they need and by the potential for synergy between different organisms. Comparisons of culturable and total microscopic cell counts from diverse habitats have demonstrated the inadequacy of the culture-dependent approach to analyse microbial community composition (summarized in Amann et al., 1995). Therefore, other tools are required to supplement the conventional microbiological techniques. The introduction of molecular techniques in microbial ecology, including those that use the gene sequences of the small subunit ribosomal RNA as a molecular marker for identification of microorganisms, has changed our perception of the diversity of microbial communities. The genes encoding small subunit ribosomal RNAs reflect the evolutionary relationship of microorganisms
METHODS IN MICROBIOLOGY, VOLUME 30 ISBN 0 12 521530-4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any form reserved
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(Woese, 1987) and the sequences of these genes allow one to group and identify microorganisms. Despite some uncertainties about the p h y l o g e n y inferred from rRNA (e.g. the rooting of the different domains) which have emerged as a result of whole-genome sequencing studies and the use of alternative molecular markers (Pennisi, 1998; Doolittle and Logsdon, 1998), the 16S rRNA approach remains the standard marker (Ludwig and Schleifer, 1999). Giovannoni and co-workers (1990), for instance, used a cultivation-independent approach consisting of PCR amplification, cloning and sequencing of 16S rRNA gene fragments to characterize the composition of Sargasso Sea bacterioplankton. The sequences obtained represented u n k n o w n 16S rRNA genes of heretofore uncultivated bacteria, and confirmed the limitations of cultivation-dependent approaches. Similar differences between culture-dependent and molecular approaches were observed by Ward and colleagues for a hot spring cyanobacterial mat c o m m u n i t y (Ward et al., 1990) and have often been reported from microbial ecology studies (for a review see Muyzer, 1998). To get a better insight into the temporal dynamics or spatial variation of microbial communities, microbial ecosystems need to be studied over longer periods of time (e.g. days to years) or samples from m a n y different locations have to be analysed. Although successful, the application of cloning and sequencing of 16S rRNA genes is too laborious and time consuming to analyse a large n u m b e r of samples, even with the recent progress in sequencing technology. Genetic fingerprinting techniques, however, are excellently suited to comparison of large numbers of samples. Genetic fingerprinting of microbial communities provides banding patterns or profiles that reflect the genetic diversity of the community. Denaturing gradient gel electrophoresis (DGGE) of PCRamplified gene fragments is one of the genetic fingerprinting techniques used in microbial ecology (Muyzer, 2000). In DGGE similar-sized DNA
Figure 22.1. Flow diagracn of PCR-DGGE analysis of microbial communities. Tile different steps are discussed in detail in the text. Briefl'~;bacteria are collected on filters, their nucleic acids are extracted and used as template in tile PCR. The mixture of PCR products is analysed by DGGE. Community profiles can be further analvsed with statistical methods, such as UPGMA and MDS (see Figure 22.3 for an example). To identify the community rnembers, bands are excised from the denaturing gradient gels, re-amplified and sequenced. The sequence data are used for phylogenetic analysis, or can be used for the design of specific probes to detect bacterial cells iJ~ situ (see Chapter 11 this volume). The gel shows temporal shifts in the bacterial diversity of mesocosm samples which are reflected in different community profiles. The time interval between the samplings were: 2 days between samples run on lanes 1 and 2, and 3 days between samples of lanes 2 and 3 (total time between samples 1 and 3:5 days). Lane H shows a marker composed of PCR roducts from five different DNAs (see section on DGGE standards). Sequences determined from the DGGE bands are shown in bold type in the tree. The phylogenetic tree has been created with the special parsimony tool implemented in the software program ARB (Ludwig et al., 1998, Strunk and Ludwig, 1998), which allows reliable positioning of the partial sequence data in a tree derived from complete sequences, without affecting the topology of the tree. 427
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fragments are separated in a gradient of DNA denaturants according to differences in sequence. A variant of DGGE, temperature gradient gel electrophoresis (TGGE), makes use of a temperature gradient to separate gene fragments. DGGE is relatively easy to perform and is especially suited to the analysis of multiple samples. Since its introduction into microbial ecology by Muyzer et al. (1993) it has been adapted in many laboratories as a convenient tool for the assessment of microbial diversity in natural samples. A general overview of PCR-DGGE fingerprinting of microbial communities is shown in Figure 22.1.
Principle of DGGE separation Amplification of DNA extracted from mixed microbial communities with primers specific for 16S rRNA gene fragments of bacteria result in mixtures of PCR products. Because these products all have the same size, they cannot be separated from each other by agarose gel electrophoresis. However, sequence variations between different bacterial rRNAs bring about different melting properties of these DNA molecules, and separation can be achieved in polyacrylamide gels containing a gradient of DNA denaturants, such as a mixture of urea and formamide. PCR products enter the gel as double-stranded molecules; as they proceed through the gel, the denaturing conditions gradually become stronger. PCR products with different sequences therefore start melting at different positions (i.e. at different denaturant concentrations) in the gel. Melting proceeds in so-called 'melting domains'. Once a domain with the lowest melting temperature reaches its melting temperature at a particular position in the denaturant gradient, a transition from a double-stranded to a partially melted molecule occurs. The protruding single strands practically cause a halt of the molecule at that position. To prevent the complete dissociation of the two DNA strands, a 40-nucleotide GC-rich sequence ('GC-clamp') is attached at the 5'-end of one of the PCR primers.
Applications of PCR-DGGE in marine microbial ecology PCR-DGGE fingerprinting is a tool for monitoring variations in microbial genetic diversity, providing a minimum estimate of the richness of predominant community members. Furthermore, DGGE facilitates the identification of individual populations by hybridization analysis of patterns with specific probes, or by sequence analysis of individual bands. PCR-DGGE has been used to investigate the diversity of microbial communities, to determine the spatial and temporal variability of bacterial populations, and to monitor community behaviour after natural or induced environmental perturbations. It has been used to study communities in various habitats, such as soil, sediments, water column, hydrothermal vents, microbial mats, mesocosms, or sewage treatment plants. Here we will give some examples of the application of PCR-DGGE 428
in marine ecosystems. For a more comprehensive overview of the use of PCR-DGGE in microbial ecology the reader is referred to Muyzer (1998, 1999) and Muyzer and Smalla (1998).
DGGE to study spatial and temporal variability of bacterial populations The distribution of microbial populations in the marine water column depends on numerous factors and variables. Especially in stratified systems exhibiting strong physicochemical gradients, DGGE fingerprinting can reveal a concomitant stratification of resident microbial assemblages. Teske et al. (1996) used PCR-DGGE to study the distribution of sulphate-reducing bacteria (SRB) in a stratified Danish fjord. PCRDGGE combined with hybridization analysis showed that the presence of SRB increased at and below the chemocline. Most-probable number (MPN) counts of SRB were done in parallel and showed a similar trend for the distribution of SRB. Interestingly, DGGE patterns of PCR products obtained from cDNA after reverse transcription of RNA, representing the active populations, were different from those obtained after amplification of genomic DNA. Despite the agreement between MPN and DGGE, the hybridization of DGGE patterns with oligonucleotide probes and sequencing analysis of DGGE bands revealed that the SRB enriched in the MPN-tubes had a different phylogenetic affiliation from the SRB detected in the natural samples. The finding that SRB obtained from the MPN cultures belonged to the genera Desulfiovibrio, Desul[obulbus, and Desulfobacter, but those in the DGGE patterns of natural samples represented an independent lineage of the 8-Proteobacteria, verified the potential disagreement between culture-dependent and molecular methods due to selection of culturable types of SRB. The potential of PCR-DGGE for the analysis of large sets of samples was recognized by Ferrari and Hollibaugh (1999). They processed 100 samples from different stations in the Arctic Ocean to analyse the spatial variation in the diversity of bacterioplankton assemblages. DGGE fingerprints of the samples were subjected to image analysis and the spatial variation of the bacterioplankton assemblage was inferred by regression analysis of the similarity of densitometric curves derived from the DGGE patterns. The resulting dendrogram separated all DGGE patterns into five major clusters with minimally 80% similarity. While clustering of some samples corresponded to samples taken in a specific region of the Arctic Ocean, there was no correlation of geography and clustering of other samples. The authors noted that clustering of the majority of samples rather seemed to reflect different phases of the cruise and might therefore be confounded with temporal variation over the 44 day period of the cruise (Ferrari and Hollibaugh, 1999). The bacterioplankton assemblages of two estuaries in California, San Francisco Bay and Tomales Bay, differing markedly in a number of physical and biological factors, had been shown to differ in metabolic properties. The analysis of samples from both estuaries by PCR-DGGE
429
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supported the hypothesis that metabolic differences were reflected in a different bacterioplankton composition (Murray et al., 1996). Yet, a few bands were common in all samples, and a number of bands were detectable at different times in both estuaries, raising the question as to what extent factors such as the relative activity of the detected populations or metabolic plasticity might influence the differences in metabolic profiles (Murray et al., 1996). In another stud}; Murray and colleagues (1998) addressed spatial as well as temporal variations in bacterial community composition in the waters around Anvers Island (Antarctica). No obvious variation was detected between samples taken within one month from different points in an area of about three square nautical miles (3 and 50m depth). However, samples retrieved from several depths up to 1200 m on two occasions within seven weeks showed variations in DGGE patterns especially at depths of 500 and 1200 m, indicating compositional changes of the bacterial community. The authors argued that due to the low bacterial activity (estimated by leucine incorporation) advective mixing processes rather than bacterial growth might have caused most of the variation. Seasonal variation in bacterial community composition of the surface waters was inferred from changes in DGGE fingerprints over a period of almost 9 months at one station. Interestingly, the number of phylotypes decreased during the transition from spring to summer and increased from summer to autumn. Riemann et al. (1999) used PCR-DGGE to map the genetic diversity of bacterioplankton in the surface, mid- and deep water of the Arabian Sea during two consecutive monsoon periods and concluded that there was a high horizontal homogeneity of the microbial assemblages. Moreover, the dominant bands in DGGE profiles of the bacterial communities sampled eleven months apart, were remarkably similar, suggesting that if there was a seasonal variation in the bacterioplankton assemblage, it might be a predictable one. Predominant phylotypes were identified by cloning and sequencing of DGGE bands and were members of groups common in oceanic waters, e.g. members of the SARll-cluster and the cyanobacteria. However, it was remarkable that none of the bands corresponded to yProteobacteria or to members of the Cytophaga-Flavobacterium-Bactetvides phylum (CFB), and that 16S rRNA gene fragments similar to those of magnetotactic bacteria were retrieved. West and Scanlan (1999) investigated the genetic structure of Ptvchlorococcus communities by molecular methods in two depth profiles from the surface to around 100m water depth in the Eastern North Atlantic, to assess the distribution of high-light (HL) and low-light (LL) adapted populations. Cloning and sequencing, as well as hybridization with HL- and LL-specific gene probes of DNA amplified from different depths were performed. PCR products amplified with a cyanobacteriaspecific primer (N~bel et al., 1997) and a Prochlorococcus-specific primer were separated on DGGE. All three methods indicated a niche-partitioning of Prochlorococcus genotypes HL and LL in the water column and provide genetic support for flow cytometric observations of dim and bright Prochlorococcus populations. 430
DGGE to monitor population shifts after environmental perturbation As pointed out above, PCR-DGGE analyses can be performed with DNA as well as with RNA. While DNA-derived PCR amplified 16S rRNA gene fragments are related to the presence of different bacterial populations, analyses of rRNA-derived PCR products can give an indication of which bacterial populations contribute to the RNA pool. As the cellular concentration of ribosomal RNA is related to the (recent) activity of cells it helps in surveying changes in the activity of bacterial populations. An example of potential differences between DNA- and RNA-derived DGGE fingerprints is shown in Figure 22.2. Similarly, analysis of the genetic diversity and expression of functional genes can be performed using either DNA or mRNA. Here, PCR-DGGE analysis of DNA-derived PCR products show the genetic diversity (presence) of certain functional genes, while PCR products obtained after DNase digest and reverse transcription of mRNA show the diversity of expressed genes (Wawer et al., 1997) Rossello-Mora et al. (1999) investigated the response of the microbial community of marine sediments to amendment with cyanobacterial biomass under anaerobic conditions. Fluorescence in situ hybridization (FISH), DGGE of PCR products obtained from DNA as well as from cDNA after reverse transcription of RNA, and sequencing of 16S rDNA PCR products were used to assess changes in the microbial community composition. Concomitant changes in the activity of the community were followed by measurements of carbon mineralization, sulphate reduction, and ammonium production rates. Addition of cyanobacterial biomass resulted in marked changes in the composition. Dominant bands from RNA-derived banding patterns were affiliated with members of the CFB. FISH with probes specific for these CFB populations showed that, although sulphate reduction was the main mineralization process, members of the CFB, but not SRB showed the highest increase in abundance as detected by FISH. The authors concluded that these CFB played an important role in the anaerobic decomposition of complex organic matter and suggested that CFB might be responsible for hydrolysis of macromolecules and fermentation. Mesocosm experiments were performed by Lebaron et al. (2001) and Schiller et al. (2001) to study changes in the activity and diversity of bacterial assemblages from the Mediterranean Sea after addition of nutrients. Fluctuations in activity were recorded in parallel to variation in community composition, which was assessed by PCR-DGGE. Different phases were observed during the incubation corresponding to an initial increase of bacterial numbers, followed by an increase of heterotrophic protozoa cropping the bacterial production and a new increase of bacterial production after the peak in grazing activity (growth, grazing, and post-grazing phase, respectively). These phases were reflected by concomitant changes in DGGE-fingerprints of the bacterial assemblage. Both, nutrient addition as well as grazing of protozoa seemed to effect changes in the bacterial genetic diversity. Multidimensional scaling analysis of DGGE patterns showed that differences in the development of the bacterial communities
431
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occurred between nutrient-enriched and control mesocosms and indicated that duplicate treatments were reproducible. Sequencing of DGGE bands was used to identify several microbial populations. DGGE bands of some populations disappeared from the DGGE patterns during the grazing phase, while members of the Cytophaga-Flavobacterium-Bacteroides phylum and Ruegeria-like bacteria became especially important after the peak in grazing activity. The latter populations also dominated the RNA-
1
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432
derived D G G E -fingerprints and hence it w a s suggested that these p o p u lations escaped the grazing pressure and were i m p o r t a n t contributors to bacterial p r o d u c t i o n a n d activity in the post-grazing p h a s e of the experiment. In a multidisciplinary a p p r o a c h McCaig et al. (1999) studied the impact of fish f a r m i n g in cages on N-cycling a n d c o m m u n i t y structure of the u n d e r l y i n g sediment. Organic carbon content and a m m o n i u m concentration of the s e d i m e n t were m e a s u r e d along a transect from the fish cage to a distance of 40 m from the cage. Carbon content and a m m o n i u m concentration were m u c h higher u n d e r the fish cage than at the other s a m p l i n g sites along the transect. Furthermore, nitrification and denitrification were strongly inhibited b e n e a t h the fish cage. D G G E was used to profile the diversity of b e t a - a m m o n i u m oxidizing bacteria ([3-AOB), along a transect from u n d e r n e a t h the fish cages to 40 m from the cage, after PCR amplification of 16S r R N A gene f r a g m e n t s with p r i m e r s specific for [3-AOB ( K o w a l c h u k et al., 1997). DGGE profiles w e r e blotted onto a m e m b r a n e and h y b r i d i z e d with probes specific for subclusters of the [3-AOB to reveal the identity of [3-AOB populations. D G G E profiles of highly polluted sedim e n t s u n d e r the fish cage s h o w e d two p r o m i n e n t b a n d s that were only faintly visible in D G G E profiles of s a m p l e s f r o m 20 and 40 m from the cage. ¢-
DGGE to study archaea, eukaryotes, and viruses The e x a m p l e s described a b o v e s h o w that PCR-DGGE is increasingly being u s e d in ecological contexts to get a better u n d e r s t a n d i n g of the factors regulating bacterial c o m m u n i t y composition. Bacterial c o m m u n i t y composition, however, is not only influenced by physicochemical factors, but m a y be effected b y biotic factors such as p r i m a r y production, grazing (Jfrgens et al., 1999; Pernthaler et al., 1997; Sch~fer et al., 2000; Suzuki,
Figure 22.2. DGGE patterns of PCR-amplified 16S rRNA gene fragments obtained from bacterial DNA, indicating the presence of bacteria, or from bacterial ribosomal RNA, showing the most active populations within tile assemblage. Water samples from two different locations were analysed: sample A (lanes 1 and 2) is from surface water taken off the coast of Banyuls-sur-mer (France) in May 1997; sample B (lanes 3 and 4) is from coastal water taken near the mouth of the river RhOne (France) in April 1998. Fingerprints of the natural bacterial assemblages were obtained from DNA (lanes 1 and 3) or after DNA digestion and reverse transcription of RNA (lanes 2 and 4). Note the differences in DNA- and RNA-derived patterns in the upper part of lane 1 and 2, where some of the bands seen in the DNA-derived pattern are weaker or not represented at all in the RNA-derived patterns. This indicates that corresponding populations have a relatively low RNA content, and hence are probably less active than others. The marked difference in intensity of the band at the bottom of the profiles shown in lanes 3 and 4 indicates that a population contributes relatively less to the DNA pool, but relatively more to the RNA pool indicator of recent cellular activity. Hence, cells of this population probably have a high rRNA content, which might indicate that they are active.
433
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1999; van Hannen et al., 1999a) and viral infection (Fuhrman, 1999; van Hannen et al., 1999b). Therefore, it is of great interest to extend the molecular approach to study the genetic diversity of primary producers, grazers and viruses. Furthermore, Archaea might also play an important role in the marine system (DeLong, 1992). Other than analysing bacterial communities by PCR-DGGE, specific PCR-DGGE assays allow one to study eukaryotic microorganisms (van Hannen et al., 1998) and Archaea. Furthermore, PCR-DGGE applications for viral communities have been introduced (Scanlan and Wilson, 1999; Short and Suttle, 1999). These assays will be valuable for a more integrated study of microbial communities. Reports of unusual crenarchaeal 16S rRNA gene sequences retrieved from marine waters (DeLong, 1992, Fuhrman eta[., 1992) have triggered further studies into the importance of Archaea in the marine environment. It has been shown by oligonucleotide hybridization that archaeal rRNA may amount to a high percentage of total rRNA extracted from concentrated marine picoplankton (Murray et al., 1998), suggesting that Archaea have been largely ignored as potentially important members of marine microbial communities. Different PCR-DGGE assays have been used for the analysis of archaeal 16S rRNA gene fragments (Casamayor et al., 2000; Ovre~s et al., 1997; R611eke et al., 1998; Vetriani et al., 1999). PCR-DGGE analyses of microbial communities from meromictic lakes by Gvred~s and colleagues (1997) and by Casamayor et al. (2000) have also reported crenarchaeal sequence types related to those reported from coastal surface waters (DeLong, 1992), marine archaeoplankton communities however, have not yet been analysed by PCR-DGGE. DGGE fingerprinting is not limited to the use of 16S rRNA gene fragments, but can also be performed with functional genes (Fesefeldt and Gliesche, 1997; Wawer and Muyzer, 1995). Using primers targeting the gene encoding [NiFe] hydrogenase of Desulfovibrio species Wawer et al. (1997) were able to analyse expression of these genes in complex microbial communities by DGGE.
Eukaryotic microbial communities Although small eukaryotes such as protozoa can be identified much easier by microscopy than bacteria because of their discriminative morphological features, their identification is time consuming and can often be done by experts only. Identification of eukaryotic microbes by molecular methods can be achieved with primers developed by van Hannen et al. (1998), which amplify a 210 bp 18S rRNA gene fragment that can be separated by DGGE. Due to the limited size of the fragment, sequencing of gel bands may make identification possible at the phylum level only (van Hannen et al., 1998). However, the authors demonstrate that, using species-specific oligonucleotide probes for hybridization analysis of DGGE gels, identification at the species level is possible. Van Hannen et al. (1998) used the eukaryote-specific PCR-DGGE assay to compare the diversity of five Dutch lakes of a lagoon system. Analysis of
434
DGGE-fingerprints and environmental variables of these lakes by UPGMA resulted in similar clustering of lakes and the respective genetic fingerprints of their eukaryotic diversity.
Viral communities Viruses are a numerically important part of the microbial food web. Virusinduced mortality may contribute significantly to overall mortality of natural microbial populations (Suttle, 1994) and has therefore the potential to affect the diversity of bacterioplankton and primary producers (van Hannen et al., 1999b). Unfortunately, viruses do not contain ribosomal RNA, hence, for the study of natural virus communities other molecular markers are needed. Essential virus genes present in a large number of certain virus groups are candidates for molecular assays, and applications of PCR-DGGE assays to study diversity in virus communities have been described by Short and Suttle (1999) and Scanlan and Wilson (1999). The assay by Short and Suttle is based on primers that specifically amplify gene fragments of the DNA polymerase genes (pol) of viruses infecting microalgae (Phycodnaviridae) (Chen and Suttle, 1995; Short and Suttle, 1999). Short and Suttle suggested that similarly, the development of primers specific for DNA polymerases of cyanophages and bacteriophages should be possible. Separation of virus PCR-amplified pol-gene fragments derived from cultures was accomplished by DGGE. A preliminary analysis of natural marine Phycodnaviridae communities demonstrated that they may undergo seasonal changes, and that community composition may vary over relatively small spatial scales (Short and Suttle, 1999). Scanlan and Wilson (1999) have applied a cyanophage-specific PCRDGGE assay based on primers described by Fuller et al. (1998) which target genes encoding capsid-proteins. DGGE was used to separate fragments of PCR-amplified virus-capsid protein genes obtained from a variety of virus strains. Application of cyanophage DGGE fingerprinting holds promise for gaining more insight into the influence of cyanophages on the diversity of cyanobacterial populations. Furthermore, they might facilitate elucidation of environmental stimuli, e.g. phosphate-limitation (P-limitation) that might decide whether a lytic, pseudolysogenic or lysogenic infection is established by cyanophages. Wilson et al. (1998) induced P-limitation in a mesocosm experiment by the addition of excess nitrogen (N) at an N:P ratio of 75:1. Nutrient addition led to a large Syllechococcus bloom, which was shown to become P-limited by using an immunological marker for P-limitation in Synechococcus. Virus concentrations increased at the same time, just before the bloom collapsed after re-addition of P. The authors suggested that the P status of the Synechococcus population had important implications for the result of the host/phage interaction. They pointed out, however, that there was not sufficient data and too much variation in virus abundance estimates to establish whether or not the collapse of the Syllechococcus bloom was due to lysogenic or pseudolysogenic host/phage interaction. 435
In the above section we have shown that PCR-DGGE is a useful tool for the analysis of complex microbial communities, and is contributing to change our perception of how the diversity of microbial communities is controlled. The examples discussed emphasize that DGGE is not a standalone technique, but should be used in combination with other measurements related to the physicochemical and biotic factors that regulate the activity and diversity of microbial communities. The different steps of the PCR-DGGE approach are described in the following section. The method can also be used to study microbial communities from other ecosystems, although some modifications in sample preparation and processing may be necessary.
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PRACTICAL ASPECTS OF PCR-DGGE
Equipment Apart from standard laboratory facilities, such as a refrigerator, freezer (-20°C and -80°C), and fume hood, the basic equipment needed for the analysis of bacterial communities by PCR-DGGE consists of: • Benchtop centrifuge with refrigeration for I 5 ml tubes • Benchtop centrifuge for 0.5 and 1.5 ml tubes • Thermocycler • Water bath • Ice-bath • Vortex • Power supplies for electrophoresis systems • Agarose gel electrophoresis system • Denaturing gradient gel electrophoresis system, e.g. DCode system (Bio-Rad 170-9080; includes gradient former) or PhorU system (Ingeny) • UV-transilluminator and Polaroid camera or a fluorescence imaging system, e.g. Fluor-S Multiimager (Bio-Rad 170-770 I, Macintosh version)
Optional equipment •
Peristaltic pump (Model EP- I, Bio-Rad 73 I-8142) and gradient former (Model 385, Bio-Rad 165-2000) for casting gradient gels • Automatic D N A sequencer, e.g.ABI 310 Genetic Analyzer (Perkin Elmer)
Sampling of bacteria About 10" bacterial cells are collected from water samples by gentle filtration on hydrophilic Durapore filters (Millipore, GVWP04700, polyvinylidene fluoride membrane, 0.22 lam, 47 mm diameter). After filtration, the filters are transferred into cryovials and immediately frozen in liquid nitrogen or otherwise. Upon return to the laboratory the filters can be stored at -80°C.
436
E x t r a c t i o n of nucleic acids Several protocols have been published in the literature for extraction of nucleic acids from marine microorganisms. We routinely use the protocol described here; it represents a combined extraction of D N A and RNA from bacteria collected on Millipore GVWP filters. The protocol is also suited to extraction of DNA and RNA from Gram-positive bacteria (Rossello-Mora et al., 1999), and Archaea (Casamayor et al., 2000).
Reagents and disposables • • • • • • • • • • •
Safety glasses, lab coat and gloves 15 ml sterile disposable centrifuge tubes 1.5 ml sterile disposable microcentrifuge tubes Tube racks AE buffer (20 mM sodium acetate, I mM EDTA, pH 5.5) Phenol:chloroform:isoamylalcohol (25:24:1), pH 7 (PCI) 10% (w/v) sodium dodecyl sulphate (SDS) 3 M sodium acetate, pH 5.2 100% (v/v) ethanol 70% (v/v) ethanol Water (Sigma W4502) or TE-buffer (I 0 mM Tris, I mM EDTA, pH 8.0)
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Method 1. Pre-warm the PCI to 60°C in a water bath. Pre-cool the AE buffer on ice. Set the temperature of the centrifuge at 4°C. 2. Transfer the filter from the cryovial to a 15 ml tube with clean forceps. Keep the tube on ice. 3. Rinse the filter with 2 ml of ice-cold AE buffer. Vortex briefly, and put the tube back on ice. 4. A d d 5 ml of hot PCI and 150 pl of SDS. Incubate for 5 rain at 60°C. Vortex briefly every minute. 5. Cool the tube on ice. 6. Centrifuge at 4000g for 5 min at 4°C to separate the aqueous and organic phase. 7. Transfer the aqueous phase to a clean 15 ml tube, and add 1/10 of a v o l u m e (ca. 200 pl) of s o d i u m acetate. S. Add 5 ml of PCI. Vortex briefly, and separate the two phases as described in step 6. 9. If necessary, repeat steps 7 and 8, until no proteineous material is visible at the interface between the aqueous and organic phase. 10. Transfer the aqueous phase to a clean 15 ml tube. Add 2.5 volumes of ice-cold 100% ethanol, and incubate for at least 3 h at -20°C. 1l. Centrifuge at 4000g for 60 min at 4°C to pellet the precipitated nucleic acids. Remove the supernatant by gentle aspiration (use a fresh sterile pipette tip for each sample). 12. Rinse the DNA pellet with 1 rnl of ice-cold 70~ ethanol. 13. Centrifuge at 4000g for 5 min at 4°C. Remove the supernatant.
437
14. Dry the pellet under vacuum. 15. A d d 100 111of water (or TE buffer) and incubate overnight at 4°C. 16. Re-dissolve the pellet by gentle pipetting. Aliquot the nucleic acid solution into sterile reaction tubes and store at -80°C. Inspect 5 btl of the solution by electrophoresis in a 1% (w/v) agarose gel together with an appropriate molecular weight standard, e.g., lambda Hind IIIdigest (Stratagene 201109). Notes
• All steps involving the handling of phenol or phenol-containing solutions should be performed in a fume hood, wearing safety glasses, gloves, and a lab coat.
Purification of R N A Reagents and disposables • • • • • • •
10x DNase buffer (400 mN Tris, 60 mN MgCI2, pH 7.5) DNase I, RNase-free (10 u pl ', Pharmacia 27-0514-01) 3 N sodium acetate pH 5.2 Phenol:chloroform:isoamylalcohol (25:24:1), pH 7 (PCI) 1.5 ml sterile disposable microcentrifuge tubes Tube racks Safety glasses, lab coat and gloves
Method
1. Add the following reagents to a 1.5 ml tube: 2 121water 2 t21 10x DNase buffer 15 111nucleic acid extract 2. A d d 1 tll of DNase, and incubate for 30 min at 37°C in a water bath or thermocycler. 3. A d d 280 111 of water and 30 111 of sodium acetate. Vortex briefly. Remove the DNase enzyme by extracting with 300 111of PCI. 4. Centrifuge at 4000g for 15 min at 4°C to separate the aqueous and organic phase. 5. Transfer the aqueous phase to a clean 1.5 ml tube. 6. A d d 2.5 volumes 100% ethanol and incubate for 2 h at -20°C. 7. Centrifuge at 4000g for 1 h at 4°C. 8. Remove the supernatant by gentle aspiration. Rinse the RNA pellet with ice-cold 70% ethanol. 9. Dry the pellet under vacuum and re-dissolve in 15 111of water. 10. Use the solution directly for first strand synthesis and store the remainder at -80°C. Notes
• All steps involving the handling of phenol or phenol-containing solutions should be performed in a fume hood, wearing safety glasses, gloves, and a lab COat.
438
•
To avoid contamination with RNase enzymes all solutions should be prepared with RNase-free water, and chemicals should be molecular biology reagent grade ('RNase - - none detected').AII steps should be done wearing gloves, additionally all lab bench surfaces and pipettes should be wiped with 70% ethanol. • Using the above pipetting scheme the maximal amount of nucleic acids solution that can be used for preparation of RNA is 171ul. In our experience 10-15 lul of nucleic acid extract usually is a sufficient amount to perform RTPCR analyses. In case of nucleic acid extracts that are low in RNA content the above scheme must be scaled up (e.g. to 100 lul volumes or more).
Preparation of first strand c D N A Reagents and disposables • •
Sterile PCR reaction tubes (0.2 or 0.5 ml) Random hexanucleotides (1:50 [v/v] dilution of Boehringer 1277081 in water (SigmaW4502) ca. 40 ng lul ') • dNTP solution (2.5 mM each dNTP; prepared from ultrapure dNTP-set [100mM each dNTP; Pharmacia 27-2035-01] and PCR water [Sigma W4S02] • 5x RT buffer (250 mM Tris-HCl, pH 8.3 at 25°C, 375 mM KCl, 15 mM MgCl 2, 50 mM dithiothreitol) • MMLV-reverse-transcriptase (200 u pl '; Promega M 1701)
Method 1. A d d I pl of hexanucleotides (ca. 40 ng t.ll t) to 10 pl of R N A p r e p a r a t i o n in a microcentrifuge tube. Incubate for 10 min at 70°C in a w a t e r bath or thermocycler to d e n a t u r e the RNA. Cool on ice. 2. Centrifuge briefly to collect the liquid in the b o t t o m of the tube and add 4 pl of 5× RT buffer and 4 pl of d N T P solution. Incubate for 2 min at 37°C. 3. A d d 1 lJl (200 u) of MMLV reverse transcriptase and incubate for 1 h at 37°C 4. Incubate the tube at 95°C for 5 rain. Cool on ice. 5. Use 1-5 lal of the solution as template in the PCR (or m a k e dilutions if too m u c h PCR p r o d u c t is obtained). Store the r e m a i n d e r at -20°C. Notes • We do not determine the RNA concentration, but by following this protocol 10 lul of RNA preparation is usually sufficient for obtaining PCR products. • It is important to save some of the RNA preparation for PCR controls to check the completeness of the D N A digestion. • According to the above scheme, I lul of c D N A preparation corresponds to an initial input of 0.5 lul nucleic acid extract containing D N A and RNA.This will be important to consider for PCR controls.
439
PCR and RT-PCR for DGGE Reagents and disposables • • • • • •
Sterile PCR reaction tubes I 0x PCR reaction buffer ( 100 mM Tris-HCI, pH 9.0, 15 mM MgCI2, 500 mM KCI) Taq DNA-polymerase (5 u tJl '; Pharmacia 27-0799-02) dNTP solution (2.5 mM each dNTP; prepared as above) Primers (50 luM) (seeTable 22.1) Water (SigmaW4502)
Method Make a 10-fold serial dilution of the extracted DNA and test several dilutions in the PCR to find the best concentration of template DNA that gives a good specific product. Note that at very high dilutions, some less abundant templates in the mixture may be lost and hence not be amplified sufficiently to form a band in the DGGE analysis. 1. Prepare a master mix for the PCR reactions by adding for each reaction the following reagents: 10x PCR reaction buffer dNTPs (2.5 mM each) Forward primer (50 laM) Reverse primer (50 pM) Water Taq DNA polymerase (5 u pl ')
10 pl 10 pl
1 pl 1 pl 76.8 pl 0.2 pl
2. Vortex, and spin briefly to collect the reagents in the bottom of the tube. Dispense 99 pl to each of the PCR reaction tubes. 3. Add 1 gl of DNA (or cDNA) solution to each tube. Close the lid, and mark the tubes. Note which template and dilution was added to which tube. 4. Spin briefly to collect all the fluid in the bottom of the tube. If your thermocycler is not equipped with a heated bonnet, overlay the reactions with a drop of mineral oil (e.g. Sigma M8862). 5. Insert the tubes into the thermocycler and start the appropriate PCR program (see Table 22.2). 6. When the run has completed store the reactions at 4°C or at-20°C until further use 7. Inspect 5 pl of the PCR products by electrophoresis on 1-2% (w/v) agarose gels together with appropriate size and mass standards. Notes
• Always perform the following PCR controls: (i) without addition of D N A template (negative control), and (ii) with addition of known D N A (positive control).
440
• We prefer to use single PCR tubes with attached lids instead of strips of tubes with strips of caps.To avoid the risk of cross-contamination of samples during pipetting of template DNAs, the tubes are opened one at a time to add template D N A and immediately closed before proceeding to the next tube. • For RT-PCR, an extra control reaction is necessary to check for the completeness of D N A digestion. A volume of RNA preparation corresponding to the amount of cDNA used as template in the PCR is added to a separate PCR reaction. If this control gives a product, then the DNA digestion was not complete, and traces of genomic D N A were present. • Addition of bovine serum albumin (BSA; Sigma B6917) to the PCR reactions may help to overcome the effects of inhibitory substances present in the nucleic acid preparations, such as humic acids. Use a final concentration of up to 3 mg ml ' in PCR reactions.
Quantification of PCR products An important point to consider in comparative DGGE analysis of multiple samples is that similar amounts of PCR products should be loaded onto the denaturing gels. Faint bands visible in one lane might not be detectable in another lane. If the total amounts of PCR product applied in these two lanes differ markedly, a fair comparison between samples is not possible. We use a mass standard (Precision Molecular Mass Standard; Bio-Rad 170-8207) for quantification of PCR products by agarose gel electrophoresis. At least three lanes are loaded with undiluted, 2-fold diluted and 4-fold diluted mass standard. Using the software MULTIANALYST (Bio-Rad) the pixel density values of marker bands with varying DNA amounts (100, 70, 50, 35, 25, 20, 17.5, 12.5, 10 and 5 ng) can be used for regression analysis. Pixel densities are also read of u n k n o w n PCR product bands, as well as of the background staining which is determined at least at one representative point of the gel. The regression curve and formula derived from the marker fragments is then used to estimate the DNA concentration of the u n k n o w n samples. Notes • Samples that are out of the range of the standard curve should be avoided. instead, these samples shouid be analysed again with more or Bess sampie applied. • • •
Avoid oversaturation of portions of the gel-pictures (by overexposure). Thoroughly rinse the ethidium bromide stained gels in Milli-Q water to increase the signal-to-noise ratio. Avoid the use of loading dyes such as xylenecyanole, as they can interfere with reliable readings of fluorescence values if they overlay D N A bands.
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Troubleshooting - - PCR No PCR product
Poor product yield Too much product or by-products Product in negative control
If no product was obtained for positive control probably due to accidental omission of a vital ingredient, try again. If positive control worked well, failure of the other reactions may be related to presence of substances inhibiting Taq polymerase in nucleic acid extracts Use more template, or pool replicate reactions and concentrate by precipitation Use less template Contamination of solutions a n d / o r plastic ware with DNA. UV-resistant plastics can be decontaminated by exposure to UV-light source (e.g. cross-linker, clean bench). Use fresh aliquots of reagents, if problems persist. Prepare new stock solutions with nucleic acid-free water
Product from RNA preparation
DNA digestion was not complete. Repeat DNA digestion with more DNase a n d / o r longer incubation time a n d / o r less nucleic acid extract
Inhibition of Taq polymerase
Substances, such as humic acids maybe co-extracted with the nucleic acids and inhibit Taqpolymerase. To overcome this problem (i) further purify the nucleic acid extracts or (ii) dilute nucleic acids which will also dilute inhibitory compounds a n d / o r (iii) add BSA to PCR reactions (see notes above)
Casting and running of denaturing gradient
gels
To achieve the maximum resolution in DGGE patterns of unknown samples it is recommended that the best gradient conditions be determined. This requires running perpendicular denaturing gels with the unknown sample to define the range of denaturant concentrations that allows the best separation possible, in our experience gradients ranging from 10-20% to 70-80% denaturant-concentration (urea and formamide; UF) result in good separation of fragments obtained by PCR with primers 341F-GC and 907R, and provide a security margin for fragments melting at unexpectedly high denaturant concentrations at the same time. It is strongly recommended that time-travel experiments be run when starting DGGE analysis to check for optimal separation. For a description of casting and running perpendicular denaturing gradient gels and time-travel experiments the reader is referred to Muyzer et al. (1996), or to the manual supplied with the DGGE system Bio-Rad.
446
Preparation of reagents Formamide (de-ionized) Add 10 g of mixed bed resin (e.g. Sigma M8032) to 100 ml of f o r m a m i d e in an Erlenmeyer and stir for 30-60 min. Decant or filter (e.g. Schleicher & Schuell folded filter 595 1/2, order no. 311647) the f o r m a m i d e to separate it from the resin beads. Store the de-ionized f o r m a m i d e in volumes of 32 ml at -20°C for the preparation of the 80% denaturing gel solution.
Acrylamide/bis-acrylarnide stock solution (37.5:1; 40% w/v) Acrylamide is a powerful neurotoxin and should be handled with extreme care. To avoid exposure to acrylamide dust, we r e c o m m e n d b u y i n g r e a d y - m a d e acrylamide/bis-acrylamide stock solution (e.g. BioRad 161-0149). If you prepare the solution from acrylamide powder, wear safety glasses, gloves, a lab coat, and a dust mask. Acrylamide Bis-acrylamide Water
38.93 g 1.07 g to 100 ml
Filter tile solution (e.g. through a Schleicher & Schuell folded filter 595 1/2) and store at 4°C in a dark bottle.
DGGE acrylamide/bis-acrylamide solutions Prepare 6~7~ (w/v) acrylamide/bis-acrylamide gradient solutions according to the amounts of reagents s h o w n below. We use 6 ~ acryla m i d e / b i s - a c r y l a m i d e solutions for PCR products obtained with primers 341F-GC/907R as well as for CYA359F-GC/CYA781R (see Table 22.1). Higher concentrations of acrylamide/bis-acrylamide m a y be necessary for DGGE analysis of other 16S rRNA gene fragments (check original citations for details). Tile use of an 80% denaturing gel solution as high denaturing solution is usually sufficient for preparation of denaturing gradient gels. However, care has to be taken that bands are not lost from the analysis due to migration to higher d e n a t u r a n t concentrations than 80~/~. In this case a 100% d e n a t u r a n t acrylamide solution should be used.
Acrylamide/bis-acrylamide 50x TAE (pH 8.3) Urea (U) F o r m a m i d e (de-ionized) (F) Milli-Q water to
0% UF
80% UF
15 ml 2 ml -
15 ml 2 ml 33.6 g
-
100 ml
32
ml
100 ml
Filter through a 0.45 p m filter or a Schleicher & Schuell folded filter 595 1/2. Degas the acrylamide/bis-acrylamide solution for 15 min u n d e r vacuum, and store at 4°C in a dark bottle.
447
10% Ammonium persulphate solution Ammonium persulphate Water
1.0 g to 10 ml
Aliquot into single-use portions and store at -20°C.
TEMED TEMED is bought as a ready-to-use solution (e.g. from Fluka or Bio-Rad).
50x TAE buffer (2 M Tris, I M acetic acid, 50 mM EDTA; pH 8.3) Tris base 0.5 M EDTA, pH 8.0 Acetic acid (glacial) Water
242.0 g 100.0 ml 57.1 ml to 1000 ml
Autoclave the buffer solution for 20 min and store at room temperature.
Ix TAE running buffer Dilute 1 volume of 50x TAE-buffer with 49 volumes of Milli-Q water.
Gradient-dye-solution To visually inspect proper gradient formation, a dye solution can be added to the high denaturant solution. Bromophenolblue (0.5% w / v final) Xylenecyanole (0.5% w / v final) lx TAE buffer
0.05 g 0.05 g 10.0 ml
10x gel loading solution Glycerol (100% v/v) Bromophenolblue (0.25% w / v final) Xylenecyanole (0.25% w / v final) Water
5.0 ml 0.025 g O.O25 g 5.0 ml
Mix and store in small aliquots at room temperature.
A s s e m b l y and casting of parallel d e n a t u r i n g g r a d i e n t
gels
1. Clean the glass plates and spacers with water and soap. Rinse them with de-mineralized water.
448
2. Wipe the glass plates first with 70% ethanol and then with acetone. Use a dust-free cloth (e.g. Kimwipes). Do not wipe any plastics (e.g. spacers, combs, etc.) with acetone. 3. Wipe the spacers (1 mm thickness) with ethanol and let them dry, then sparingly smear grease (High vacuum grease; Dow Coming, Auburn, MI, USA) along one of the long edges, such that around 2 mm are covered with a thin grease-film on each face of the spacer. 4. Place the large glass plate on a clean surface, and put the spacers onto the left and right margins, such that the greased edges face the outside edge of the glass plate. 5. Put the small glass plate on top of the spacers, to form a 'sandwich'. 6. Align the spacers and the glass plates in such a way that they are flush at the bottom of the sandwich (this can be done on an even surface, or in the aligning slot of the casting stand). 7. Attach the clamps to the sandwich, tighten the clamp screws (fingertight) and put the sandwich in the casting stand, fix in casting slot by turning the levers. 8. Before proceeding to the next step make sure that the device used for casting the gradient gel is ready installed and you are familiar with the procedure described below. The work has to proceed quickly otherwise you run the risk that gel solutions will polymerize before casting is finished. A gradient former comes with the DCode system from BioRad. We use a combination of peristaltic pump (Model EP-1, Bio-Rad 731-8142) and gradient former (Model 385, Bio-Rad 165-2000) to cast gels. For detailed instruction on set-up and operation of these refer to the technical instructions of Bio-Rad. Connect the tubing of the pump with the outflow chamber of the gradient chamber. Attach an injection needle to the Luer-lock of the outlet tubing of the pump and insert the needle between the glass plates in the middle of the gel sandwich. 9. Prepare the high and low denaturant solutions for the gradient as required in disposable plastic tubes. Using 1-mm-thick spacers, 12 ml each are recommended. The gradient gel will finally be overlaid with a 0% denaturant acrylamide solution (prepare 5 ml), as otherwise the presence of the denaturants hinders the formation of good sample wells. 10. Add ammonium persulphate and TEMED to the gradient solutions. We add 60 IJ1 of ammonium persulphate and 8 111 of TEMED to each solution, directly pipette these into the solutions. Close tubes and mix thoroughly by inverting several times. 11. To inspect the gradient, add 1201~1 of gradient-dye-solution to the high denaturant solution. Close the tube and mix by inverting several times. 12. Close the connection pipe between the two chambers of the gradient chamber, make sure that the pump is not running. Pour the high denaturant gel solution into the outflow chamber of the gradient chamber. 13. To remove air bubbles in the connection pipe, slowly open the pipe by turning the lever until the air has been expelled from the pipe and a drop of high denaturant gel solution is visible on the bottom of the 449
second chamber. Then close connection pipe and pipette back all high denaturant gel solution back to the outflow chamber using a clean pipette tip. 14. Carefully add the low denaturant gel solution into the second chamber. 15. Turn on the magnetic stirrer at 250 rpm, then turn on the pump and slowly open the connection pipe, such that no extra high denaturant gel solution enters the second chamber. Cast the gel with ca. 4 ml min '. The last 1 ml of the gradient gel will not be mixed properly (due to the remains of high denaturant gel solution in the connecting pipe of the gradient chamber), hence avoid delivery of that last bit of gradient solution, as it will disturb the top of the gradient gel. 16. Remove needle from gel sandwich. Rinse gradient chamber and pump tubing with water to remove residual gel solution. 17. Clean a comb (1 mm thick) with ethanol and let it dry. 18. Add 25 pl of ammonium persulphate and 5 pl of TEMED to 5 ml of 0~7~ denaturant gel solution. Close the tube and mix. 19. Carefully overlay the gradient gel with about half of the 0% denaturant gel solution using a 1000 pl pipette. 20. Insert the comb at an angle to avoid the formation of air bubbles. Completely fill the gel sandwich with the remainder of the 0% denaturant gel solution. 21. Let the solution polymerize for at least 2 h. Notes • There are two different kinds of spacers for the DCode system, those for casting perpendicular gels, which have grooves on the inside side of the gel sandwich, and normal spacers without grooves. For normal parallel DGGE analysis we recommend the use of spacers without grooves, as they are easier to grease and provide a better safeguard against current leakage, which may cause considerable smiling of the gel. • To facilitate mixing of the gradient solutions, the gradient chamber should be placed on a magnetic stirrer and small magnetic stirring bars should be added to each chamber. Furthermore the gradient chamber should be placed at a higher level than the peristaltic pump to improve gradient formation. • To avoid too fast a polymerization of acrylamide solutions these can be kept on ice before casting.This may be especially important when temperatures in the lab rise to high levels during summer months. • Polymerized gels can be stored overnight. To avoid drying out, the comb is removed, the wells filled with water and the gel covered with cellophane.
Troubleshooting m D G G E gel casting Acrylamide solution leaves outflow chamber of gradient former, but gel solution from second chamber is not flowing into and mixing with solution in outflow chamber
450
Mostly due to air bubbles in the connection pipe between the two chambers of the gradient former
During casting, bubbles appear in the tubing between pump and needle
Mostly due to defect pump tubing, replace tubing
During casting many air bubbles are formed at the needle that get into the gradient gel between the glass plates
Mostly due to old needle, replace needle
Acrylamide solution leaks at the bottom of the glass plate assembly
Spacers and glass plates are not flush at the bottom
Gel does not polymerize
No or not enough TEMED a n d / or APS - - added
Gel polymerizes, but remains viscous
Make sure that proper mixture or percentage of acrylamide and bis-acrylamide was used
Running parallel D G G E gels Sample preparation After quantification of PCR products, the samples are mixed with 10x gel loading solution. The total volume of PCR product to be loaded may vary between 15 lal and 60 1J1.Using a 1-mm-thick, 16-well comb of the DCode system it is possible to load volumes up to approximately 70141. Apply the sample very slowly into the sample wells to avoid mixing with the electrophoresis buffer and to avoid overflow into the neighbouring wells. As bands tend to focus in DGGE there is no need to apply equal sample volumes. Alternatively, PCR products of low concentration can be ethanol precipitated and be re-dissolved in smaller volumes.
DGGE standards Sometimes more than 20 samples are to be compared on denaturing gradient gels, exceeding the number of wells formed with the 20-well comb, hence multiple gels are needed. Denaturing gradient gels, however, show some degree of gel-to-gel variation, caused by differences in the gradient. Therefore, it is recommended that use be made of a marker standard on tile gels that is composed of fragments halting at a range of denaturant concentrations. Such a marker facilitates gel-to-gel comparison, the marker we use routinely (see lane H of gel shown in Figure 22.1) is composed of five different fragments derived from chloroplast 16S rDNA of a Nitzschia sp., two cloned 16S rRNA genes obtained from an earlier study (SchSfer et al., 2000), and two commercially available genomic DNAs of Clostridilun pelqfringens (Sigma D1760) and Micrococcus lysodeikticus (Sigma D8259). Bands halting at high denaturant concentrations can be used to normalize the migration length of individual bands which may vary between gels (Ferrari and Hollibaugh, 1999).
451
DNA
amounts
t o load
There is no general rule for the amount of PCR product to apply on denaturing gels, since the optimal amount will depend on the number of different sequence types (i.e. bands) in a given sample, as well as the relative contribution of the bands to the total PCR product (i.e. the relative intensity of particular bands). For instance, loading 500 ng of PCR product in a situation where the fluorescence intensity is equally distributed over five different bands will be different from samples showing 30 to 40 different bands. The absolute DNA amount to be loaded should therefore be tested empirically. Typically, we use about 500 ng (range 300-600 ng) PCR product for the analysis of marine bacterioplankton communities obtained by amplification with primers 341F-GC/907R. In our experience, using around 1 pg often leads to high background and overloading of individual dominant bands, potentially obscuring some other, fainter bands. Ferrari and Hollibaugh (1999) reported that about 1 12g was the optimal amount to use, however, they often observed multiple bands for single organism templates, which may have been an effect of overloading DGGE lanes rather than representing sequence heterogeneity of multiple rrll operons. For analysis of oxygenic phototrophic communities N6bel and colleagues (1999) used around 500 ng. 1. Fill the electrophoresis tank with approximately 71 of lx TAE buffer. 2. Insert the core. Attach the gel to the core, attach a buffer dam at the other site. The buffer dam can be made of a large and small glass plate without spacers and held together by the sandwich clamps. 3. Carefully place the lid (i.e. the electrophoresis/temperature control module) on the electrophoresis tank. Take care that the end of the stirring bar comes in its proper position. 4. Switch on the DCode system with the on/off button on the electrophoresis/temperature control module. Switch on the buffer recirculation pump and the heating element. Set the temperature to 60°C and set the ramp rate to 0. The buffer will reach the temperature in about lh. 5. Prepare the samples by adding between 5 and 10 121 of gel loading solution. Mix the samples and spin briefly. 6. Remove the comb slowly, when the acrylamide gel is polymerized. 7. When the buffer has reached 60°C, switch off the electrophoresis unit, wait at least 15 s before removing the lid, and place the lid on the lid stand. 8. Take out the core, pre-wet the sandwich clamps of the gel sandwich and attach to core. Replace the core in the electrophoresis tank. 9. Take a 25 ml syringe, pull up the buffer from the electrophoresis tank, attach a needle and rinse the wells of the denaturing gel to remove traces of non-polymerized acrylamide. 10. Load the samples into the wells with a 50121 Hamilton syringe. Thoroughly rinse the syringe with electrophoresis buffer between the different samples. 11. Put the lid on the buffer tank, turn on electrophoresis unit and connect the cords to the power supply. 452
12. Run the gel at constant voltage of 10 V for 10 min while the temperature is brought back to 60°C. 13. If some samples cannot be loaded completely due to too large a sample volume, switch off p o w e r unit and electrophoresis unit, and repeat steps 10 and 11. 14. Run the gel at a constant voltage of 100V for 18h. The amperage should be around 35 mA. 15. After 18 h, turn off the p o w e r supply and the electrophoresis unit. Wait at least 15 s before removing the lid. Take out the core and detach the gel sandwich. 16. Remove carefully one of the glass plates as well as the spacers. Stain the gel on the glass plate with ethidium bromide solution for 30 rain (ethidium bromide 0.5 lag ml ' in distilled water). 17. Rinse the gel for 20 to 30 min in distilled water. 18. Transfer the gel to a UV-transilluminator and p h o t o g r a p h with a Polaroid camera or preferably use a gel documentation system equipped with a CCD camera and coupled to a computer (e.g. Fluor-S Multiimager, Bio-Rad). Take several photos of the gel with varying exposure times (optimal, underexposed, overexposed). Underexposed photographs may help to define very intense bands, while overexposed photographs m a y help to identify additional faint bands. Notes
• Avoid powdered gloves as they may leave a background on the gel. •
•
DGGE gels can also be stained with Sybr Green (Muyzer et al., 1998) or Gelstar (Moeseneder et al., 1999). Specific filters might be necessary to optimize the acquisition of gel images. Denaturing gels can also be stained with silver. However, this might be disadvantageous for further re-amplification and sequencing of excised bands. Gels can be easily transferred into the Fluor-S Multiimager (Bio-Rad) using a large gels-coop (Sigma G7152).Avoid scratches in the scoop as this will show
in gel images. •
In most cases, DGGE gels are I mm thin and therefore difficult to handle. However, gels can be transferred easily from UV-tables back to glass plates or moved to a blotting device or another UV-table using Whatman filter paper. Cut a piece to match the size of the gel and carefully put it on top of the gel, avoiding bubbles. Carefully lift the filter paper, make sure the gel remains attached, and put down on a glass plate/UV-table/blotting stack. Soak the filter paper completely with water (or buffer when moving to a blotting stack) and the filter paper will come off easily.
A N A L Y S I S OF D G G E P A T T E R N S DGGE patterns from mixed microbial communities m a y be very complex. Different kinds of information can be extracted from DGGE patterns, i.e. the number, position (absence or presence of particular bands) and relative intensity of bands. Furthermore, the nucleotide sequence of bands
453
can be determined. Information extracted from DGGE patterns can be subject to numerical analysis to determine the extent of variation between DGGE patterns of different samples and thus help in the interpretation of DGGE analyses. A prerequisite for comparative analysis of DGGE patterns is that similar amounts of PCR products were applied on the gel. Figure 22.3 schematically shows the steps in numerical analysis of DGGE patterns. Deciding which features of gels represent bands and which do not is of pivotal importance. DGGE patterns can be analysed with band-finding algorithms after digitization of gel photographs. Ferrari and Hollibaugh (1999), however, noted that visual inspection of gel patterns provides the most sensitive way. This agrees with our experience, although subjective assessment cannot be ruled out with visual inspection, and analysis might vary between persons. Fragments of the 16S rDNA from different microorganisms may show varying degrees of sharpness as DGGE bands, some may focus very well, whereas others remain somewhat fuzzy. These are probably intrinsic features of the melting behaviour of different nucleic acid sequences. To remain as objective as possible, all features that look like a band should be scored as such. The basic assumption in DGGE analysis is that each band in a DGGE fingerprint corresponds to a unique type of 16S rRNA gene. Yet, there are some circumstances that prompt one to think of this in relative terms (see Limitations of the PCR-DGGE approach).
Binary matrices A first step in the analysis of DGGE patterns by statistical methods, such as unweighted pair-wise grouping with mathematical averages (UPGMA) and multidimensional scaling (MDS) is to set up a binary matrix that is representative of the bands occurring in a set of DGGE patterns. The presence or absence of DGGE bands in a sample is scored as present (1) or absent (0), relative to the DGGE bands detectable in all samples of a set of DGGE patterns.
Unweighted pair-wise grouping with mathematical averages (UPGMA) UPGMA is a clustering method for binary data whereby pair-wise similarities of DGGE patterns are used to infer a dendrogram that depicts these distances in graphical form. For UPGMA analysis of DGGE patterns a binary matrix is translated into a distance matrix representing the similarities of the DGGE patterns using a similarity coefficient. Different similarity coefficients have been used by several authors. The Dice coefficient used for cluster analysis of data from restriction fragment length polymorphism (RFLP) of 16S rRNA genes (Heyndrickx et al., 1996) and ribopatterns of bacterial strains (Vachee et al., 1997) is identical to the Sorensen coefficients used by Murray et al. (1998) for calculation of pairwise similarities and the Nei and Li coefficient used by van Hannen et al. 454
binary m a t r i x
DGGE patterns ~ A ~ B R C ~ D ~ E ~ i
i
I
I
i i i
I
I I
I
I
i
i
i
i
i
i
A
B
C
D
E
1 1 0 0 1 1 0 1 1 1 1 0 0 1 1 0 1 0 1
1 1 0 0 1 1 0 1 1 1 0 0 0 1 1 0 1 0 1
0 I 0 1 0 1 1 1 1 1 0 1 I 1 I 0 0 0 0
0 1 1 1 0 ! 1 1 0 1 0 ! ! 1 0 t 0 0 0
0 0 1 1 0 1 1 1 0 1 0 1 1 1 0 1 0 0 1
i
i
l
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
I
i
I
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i
i i
i
iiiii
0 0 ! 1 0 0 1 1 0 1 0 1 1 1 0 1 0 0 1
d i s t a n c e matrix ( J a c c a r d c o e f f i c i e n t ) A A
B
C
D
E
100
B
91.7
100
C
43.8
46.7
100
D
27.8
29.4
69.2
E
27.8
29.4
57.1
83.3
100
F
21.1
22.2
46.7
69.2
83.3
100 100
Multi-dimensional scaling
UPGMA 031
0 40
0 60
O 80
I 00
I
I
I
I
I
I
I
A
O4
g
•F
% [B
.i 09 C
"E •D
E C
F
-
-
I
E
_~ -2 D
I
t
I
-1
O
1
Dimension 1
Figure 22.3. Schematic example of statistical analysis of DGGE patterns. Briefly, the presence (1) and absence (0) of DGGE bands in different samples are scored in a binary matrix. The binary matrix is translated into a distance matrix using a similar(ty coefficient (e.g. Jaccard coefficient) that is used for UPGMA or MDS.
455
(1998) and Lebaron et al. (1999) for cluster analysis of DGGE patterns. Other authors (Curtis and Craine, 1998; Ferrari and Hollibaugh, 1999; Liu et al., 1997) have used the Jaccard coefficient (Jaccard, 1908) for clustering of fingerprint patterns (T-RFLP and DGGE). This coefficient has also been used in the schematic example depicted in Figure 22.3. Both, the Jaccard and the Dice coefficients seem to be appropriate since they do not consider double absence of bands in the calculation of the pairwise similarity, and thereby avoid spuriously high similarity values in pairwise similarities of samples (i.e. DGGE patterns of two lanes in a DGGE gel) with high numbers of double-absent bands. The Jaccard similarity is calculated according to the formula: Sp.~.~,.~t = N , . / ( N , + NI~ - N~,)
where N,~ is the number of bands common to both samples (patterns) and N~ and N~ represent the total number of bands in sample A and B, respectively. The formula for the Dice coefficient as shown in Heyndrickx et al. (1996) is: S,,,,,. = 2 N . , , ~ / ( N A + N , )
where the designations of the terms are the same as given for the Jaccard coefficient. The distance matrix is further analysed by UPGMA (for examples see Lebaron et al., 1999; van Hannen et al., 1998).
Multidimensional scaling analysis (MDS) MDS is a powerful data-reduction technique that may aid in the interpretation of large sets of complex DGGE patterns. Van Hannen et al. (1999a) were the first to use this statistical method in conjunction with DGGE fingerprinting in their study on the influence of predation on the genetic diversity of a microbial community. Sch~ifer and colleagues (2001) analysed by DGGE the development of Mediterranean bacterioplankton in nutrient-enriched mesocosms. Here, MDS not only served to show deviations between control and treatment mesocosms, but also confirmed the reproducibility of duplicate mesocosms. For MDS analyses the information of the DGGE patterns is again represented as a 0/1 binary matrix, which is used to derive a distance matrix, using the Dice or Jaccard coefficient (the Jaccard coefficient is for instance implemented in the statistics software SYSTAT 7.0). MDS reduces a complex DGGE pattern to a point in a two-dimensional space (when restricted to two dimensions). When, for instance, the development of a microbial community is studied during time by DGGE, the patterns can be analysed by MDS. Connecting the dots representing consecutive samples by lines, the development of the banding patterns can be visualized (for an example see van Hannen et al., 1999a).
456
Densitometric analysis ~ relative fluorescence of D G G E bands DGGE data may also be amenable to quantitative analysis. For this, the relative fluorescence (staining intensity) of DGGE bands has to be measured. This can usually be achieved using software such as NIHimage by plotting the pixel density along the DGGE profile. This results in a peak pattern of which individual peaks and the baseline have to be defined. Subsequently relative fluorescence values can be obtained for individual bands.
Diversity indices DGGE-derived values of genetic richness and abundance (defined as relative fluorescence of DGGE bands) can be used to calculate diversity indices. Ill a study of hyper-saline microbial mat communities, N/ibel and colleagues compared the diversity of oxygenic phototrophic microorganisms in mat samples from different sites (N/ibel et al., 1999). Using a specific PCR (Ni_ibel et al., 1997) they amplified 16S rRNA gene fragments of oxygenic phototrophs and separated them by DGGE. Different samples were compared according to the number of DGGE bands detectable (i.e. genetic richness), and their relative staining intensity (i.e. evenness). Using these PCR-DGGE-defined richness and evenness values, a Shannon-Weaver diversity index could be calculated which was compared to two other cultivation independently derived diversity estimates. It is important to note that the PCR conditions have to be adjusted such that the PCR does not reach the plateau phase. Furthermore, using bacterial/universal primers with complex communities might not result in valid diversity estimates due to complex DGGE patterns.
Identification of c o m m u n i t y m e m b e r s Apart from facilitating the comparison of larger numbers of samples, DGGE fingerprinting also makes possible tile identification of predominant community members. Two approaches have been applied successfully. The first is hybridization analysis of blotted denaturing gradient gels with oligonucleotide (e.g. Brinkhoff and Muyzer, 1997; Muyzer et al., 1998) or polynucleotide probes (Heuer eta[., 1999). The second is sequencing of excised denaturing bands. The latter approach is, however, more straightforward than hybridization analysis and also more universal, because only few of the 'group-specific' target sites (Snaidr et al., 1997) lie within the fragment of the 16S rRNA encoding gene used for DGGE analysis of mixed bacterial communities.
Excision of bands and re-amplification After documentation of the denaturing gel, make a printout of the gel and mark all bands that are to be excised and sequenced. Assign each band a number and label a corresponding number of 0.5 ml reaction tubes, accordingly. 457
1. Transfer the gel to a UV-table and set the UV-table to ' p r e p a r a t i v e ' (or 'low') instead of 'analytical' (or 'high') mode. 2. Wipe a scalpel blade with ethanol a n d switch on UV-table, cut out b a n d of interest and pick it u p with the blade or with forceps. 3. I m m e d i a t e l y switch off the UV-source to m i n i m i z e the d a m a g e to the D N A b a n d s in the gel. 4. Transfer the gel piece to the labelled tube. 5. Continue excising b a n d s as described in steps 2-4, until all b a n d s have been excised.
1
2
--
3
4
5
6
7
Iii
W m
Figure 22.4. DGGE of re-amplified bands that were excised from a denaturing gel for sequence analysis (lanes 1 and 2, and lanes 4 to 6). The re-amps were run side by side with PCR products of the original samples (lanes 3 and 7) to verify that (i) the re-amplified products were single bands, and (ii) correspond to the excised band in the original pattern. Sometimes re-amplified products might consist of more than one band (e.g. lanes 5 and 6). In such cases the band should be excised and re-amplified again. Alternatively, such PCR products can be cloned to isolate the band of interest.
458
6. Rinse the bands by adding 200 t~1 of nucleic acid-free water, spin d o w n contents of tubes and incubate at room temperature for 1-2 h. 7. Remove the water by gentle aspiration (use a clean sterile tip for each band). 8. Add 25-50 pl of nucleic acid-free water, spin down, and incubate at 4°C overnight. 9. Use water from the supernatant as template for re-amplification with the same primers as for the PCR for DGGE, store the remainder at -20°C. 10. Check the PCR product from the re-amplification alongside the original DGGE pattern to make sure it is the proper band and to see if it is a single band (see Figure 22.4). Notes
• Ethidium bromide is a powerful mutagen. AIways wear at least one pair of protective gloves. • Protect yourself against exposure to UV radiation by wearing a UV-filtering face-shield. Shield arms/wrists by taping the ends of the lab coat sleeves tight around the wrists with tape. • UV-light will also damage the DNA that is to be re-amplified. Therefore, excision should proceed as quickly as possible and UV-exposure has to be kept as short as possible.This can be achieved by switching off the UV-source as soon as a band has been excised and only turning it on when you are ready for excision of the next band. • Avoid scratching the surface of the UV-table, excise bands by pressing scalpel blade carefully through the gel rather than cutting with it.
Cycle-sequencing of PCR-products Reagents and disposables PCR reaction tubes (single tubes not strips) ABI PRISM® BigDye TM Terminator Cycle Sequencing Kit (Perl
Method After inspection of re-amplified DGGE bands by DGGE the PCR products to be sequenced are purified using the Qiaquick PCR purification kit (Qiagen 28106) according to the manufacturer's instructions and the concentration of the purified products is estimated as described above. The pipetting scheme given below is based on the BigDye cycle sequencing ready reaction kit of Perkin Elmer (Perkin Elmer 4303149). It deviates from the recommendations of Perkin Elmer in that just a quarter of the r e c o m m e n d e d a m o u n t of ready reaction mix and extra buffer (BDT buffer) is used (ready reaction mix includes buffer, dNTPs, fluorescently labelled ddNTPs, Amplitaq).
459
1. For each reaction prepare in a 0.2 ml PCR tube: BDT 5× buffer Ready reaction mix DNA Primer (5 pM) Water
2.
3. 4.
5. 6.
4 pl 2 pl ca. 50 ng 0.8 pl to final volume of 20 pl
For multiple reactions with the same sequencing primer prepare a master mix consisting of ready reaction mix, 5x BDT buffer, water and primer, dispense into 0.2 ml tubes and add water and template accordingly. Note that this is the recipe for a 1/4 reaction. Briefly spin to collect the reagents in the bottom of the tube. Perform cycle sequencing: 25 cycles, each consisting of a rapid thermal ramp to 96°C - - hold for 10 s, rapid thermal ramp to 50°C - - hold for 5 s, rapid thermal ramp to 60°C - hold for 4 rain. After the final cycle include a rapid thermal ramp to 4°C and hold temperature (it might be necessary to adjust annealing temperature for particular primer sequences). Store the reactions at 4°C, until purification. Purify cycle sequencing products (e.g. by isopropanol precipitation, as recommended by Perkin Elmer) and finally re-suspend the pellets in 13 t.11of Template suppression reagent (Perkin Elmer). Denature samples for 3 min at 95°C in the thermocycler, then place on ice. Spin down briefly, transfer to ABI sample tubes and load on sequencer.
Comparative sequence analysis Analysis of sequences can be carried out using a number of different software programs and database services on the Internet, such as the Ribosomal Database Project (RDP; Maidak et al., 1999). We routinely use ARB, which is a comprehensive tool for comparative analysis of sequence data (Ludwig et aI., 1998; Strunk and Ludwig, 1998).
Reproducibility and sensitivity of PCR-DGGE The PCR-DGGE approach to study microbial communities is a highly reproducible and a consistently performing fingerprinting technique (Ferrari and Hollibaugh, 1999; Murray et al., 1996; Muyzer, 1999; f~vreas et al., 1997; Riemann et al., 1999) and it has been suggested that even if biases affect the fingerprints generated by PCR-DGGE these biases are of a constant nature and thus changes in banding patterns do reflect variations in microbial community composition (f~vre~s et aL, 1997; Riemann et al., 1999). Different studies have demonstrated the sensitivity of the PCR-DGGE approach. Muyzer et al. (1993) found that in a mixture of different template DNAs the detection threshold of dilute templates by PCR-DGGE 460
was around 1% of the total DNA. Similar values have been observed by others. Murray noted that in artificial mixtures template concentrations of 1.6% could be detected easily (Murray et al., 1996). In a study of the bacterioplankton of two Spanish lakes (Lakes Ciso and Vilar), Casamayor and colleagues (2000) could test the sensitivity of the method by comparing the identity of detected bands to counts of microscopically identifiable populations and also detected populations present at a density of around 1% of total cell counts (Casamayor et al., 2000). A new fingerprinting method introduced recently by Liu et al. (1997) into microbial ecology is terminal restriction fragment length polymorphism (T-RFLP). Marsh et al. (1998) analysed the eukaryotic microbial community of activated sludge samples and found that T-RFLP detected about twice as much phylotypes as DGGE. However, neither the primer sequences or specificity were given, nor did they describe their DGGE assay. A comparison of DGGE and T-RFLP was also made by Moeseneder et al. (1999) who analysed complex marine bacterial communities with optimized protocols for both methods. Although, T-RFLP detected a higher number of phylotypes (range 33 to 44, depending on restriction enzyme used) than did DGGE (range 28 to 35) the differences were not large.
L I M I T A T I O N S OF T H E P C R - D G G E A P P R O A C H As with all other molecular biological applications that rely on PCR as an initial step, DGGE fingerprinting is potentially afflicted with PCR inherent biases. These PCR biases have been described in detail elsewhere (yon Wintzingerode et al., 1997). The same is true for biases afflicted with unequal lysis of bacterial cells and efficiency to obtain a nucleic acid preparation representative of the community composition. Specific limitations of DGGE are that only short sequence fragments can be used (up to ca. 500 bp), thereby limiting the amount of sequence information for subsequent identification by comparative sequence analysis. Furthermore, the resolution of different sequences is not always accomplished and assignment of features as bands or not bands can also be difficult in some cases. Other biases also lead to artefacts with other molecular approaches. Bacteria may harbour more than one copy of the 16S rRNA encoding gene, with heterogeneous sequences, giving rise to more than one band on DGGE (N/ibel et al., 1996). Furthermore, dissimilar sequences may co-migrate to the same position in a DGGE gradient (Buchholz Cleven et al., 1997; Fesefeldt and Gliesche, 1997; Kowalchuk et al., 1997; Rossello-Mora et al., 1999), causing a band to be a mixture of more than one sequence and preventing recovery of a clean sequence after re-amplification. These problems potentially concern the reliable estimation of the number of different phylotypes (i.e. richness) by all genetic fingerprinting methods. Furthermore, artificial bands may be due to heteroduplex molecules (Ferris and Ward, 1997), which may form between single strands of two similar, but not identical DNA molecules. However, potential extra bands should not interfere with conclusions based on the comparison of patterns from different samples. 461
Murray et al. (1996) noted that heteroduplex formation did not significantly interfere with DGGE analysis of complex communities. Molecular approaches to investigate microbial community composition are not free of biases either and attempts to cultivate bacterial strains representative of populations important in natural communities should not be neglected, since such strains have to be investigated to get new insights into the activities of marine microbes. To reduce misinterpretation of results due to biases or limitations of the techniques used it may be of advantage if other molecular, microbiological and geochemical measurements are made at the same time.
CONCLUSIONS Numerous studies in microbial ecology have used PCR-DGGE fingerprinting for the analysis of microbial community composition up to now. It has been shown by several studies that the approach is reproducible and sensitive. These aspects as well as the straightforwardness of DGGE will probably attract even more scientists to adapt this relatively inexpensive technique as a new tool in their study of microbial ecology in the future. While other new fingerprinting techniques, such as automated T-RFLP, might be more sensitive, identification of predominant community members still requires cloning and sequencing of PCR products. A potential future development in PCR-DGGE fingerprinting might be to use fluorescently labelled PCR primers, which might (i) make staining of gels unnecessary and (ii) make it possible to add intra-lane standards with a different fluorochrome, facilitating gel-to-gel comparisons.
Acknowledgements We are grateful to many colleagues who have developed, adapted and used PCR-DGGE analyses in many ways, and we are looking forward to new developments and applications of this technique that will expand and maybe change our perception of the organization of microbial communities in the future. This work was supported by the European Union research programme 'Preserving the Ecosystem' under CHABADA project contract MAS3CT96-0047, and by the Max-Planck-Gesellschaft, Munich. It is ELOISE contribution No. 134.
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23 Measurement of UVB-induced D N A Damage in Marine Planktonic Communities W a d e H Jeffrey' and David L Mitchell 2 'Center for Environmental Diagnostics and Bioremediation, University of West Florida, Pensacola, FL 32514, USA; 2The University of Texas M.D. Anderson Cancer Center, Department of Carcinogenesis, Science Park - Research Division, Smithville, TX 78957, USA
CONTENTS
Introduction Principle and methodology Applications Data analysis Conclusions and future directions
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INTRODUCTION Ultraviolet radiation (UVR) has been recognized for many years as a potential stress for organisms in a variety of environments (Worrest at al., 1978; 1981; Calkins, 1982) and as a factor in biogeochemical cycling (Zepp at al., 1995). Direct biological effects of UVR result from absorption of specific wavelengths of light by specific macromolecules and the dissipation of the absorbed energy via photochemical reactions (Mitchell, 1995). Cellular targets of UVR include nucleic acids, proteins, membrane lipids, the cytoskeleton, and photosystem 1i (Vincent and Roy, 1993; Mitchell, 1995). Dimerizations between adjacent pyrimidine bases are the most prevalent photoreactions resulting from the direct action of UVR on DNA (Mitchell, 1995). The two major photoproducts are the cyclobutyl pyrimidine dimer (CPD) and the pyrimidine(6-4)pyrimidinone photoproduct [(6-4)photoproduct; Figure 23.1], which is converted to its valence photoisomer, the Dewar pyrimidinone, by absorption of UVB light between 310 and 320 nm (Mitchell, 1995). Formation and structural differences of the (6-4)/Dewar photoproducts and CPDs are significant and determine their different molecular and biological effects (Mitchell and Nairn, 1989). Both CPDs and (6-4) photoproducts can inhibit DNA synthesis and gene transcription, but because of structural differences, the (6-4) photoproduct is 300-fold more efficient at blocking the progression of DNA polymerase
METHODS IN MICROBIOLOGY, VOLUME 30 ISBN I1-12-521530 4
Copyright © 2001 Academic Press Ltd All rights of reproduction m anv form reserved
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Figure 23.1. DNA photoproduct formation from ultraviolet radiation. A 'normal' pair of adjacent thymine nucleotides (TpT) are converted to a DNA photoproduct by the energy of UVB radiation which creates new bonds between the adjacent bases.
than the CPD. Although induced at lower frequencies, the (6-4) photoproduct and Dewar isomer may be responsible for many of the lethal effects of UV-B radiation. Although most research examining the effects of UVR on marine microbial communities has been directed at phytoplankton and primary production, it is now apparent that all microbial trophic levels must be considered when investigating the ecological impact of UVR. The importance of bacterioplankton in oceanic processes has become widely recognized. Bacteria have been found to play a vital role in carbon cycling, transferring significant amounts of material to higher organisms. Numerous studies have shown that bacterial production is of the same order of magnitude as primary production. Although the data are quite variable and depend on location and season (Fuhrman and Azam, 1980; Hansen at al., 1983; Sullivan et al., 1990; Cota at al., 1990), bacterial production values often equal 50% of the primary production. Bacteria have been found to account for up to 90~ of the cellular DNA in oceanic environments (Paul and Carlson, 1984; Paul et al., 1985; Coffin et al., 1990) and the role of bacteria in elemental and nutrient cycling has received extensive study (Falkowski and Woodham, 1992). Bacteria, in concert with their predators (viruses and eukaryotes), play critical roles in nutrient recycling in the water column. It is plausible that reduced bacterial nutrient cycling resulting from UVB injury may result in diminished primary production. Likewise, it might be expected that a decrease in phytoplankton production may result in a decline in bacterial 470
production which may be compounded by direct UVB effects on bacterioplankton. UVR may directly impact viruses, bacteria, phytoplankton or zooplankton via direct DNA damage and reduced rates of production. Direct effects on one trophic group mav result in an indirect impact on others. The effects of UVR on marine bacterioplankton has been most often investigated by using radiolabeled precursor molecules such as ~Hthymidine (TdR; Fuhrman and Azam, 1982) and ~H- or '~C-leucine (Leu; Chin-Leo and Kirchman, 1988; Simon and Azam, 1989). Broad-band cutoff filters (e.g. Mylar 500D, UF-3 plexiglass) have then been used to selectively exclude UVB or UVB+UVA. The rate of incorporation of radiolabeled substrate is compared among treatments and compared relative to a control sample incubated in the dark. Using this method, Herndl et al. (1993) demonstrated a 48% inhibition of TdR incorporation, relative to a dark control, in surface water samples taken from the northern Adriatic Sea after 4 h incubations. Aas et al. (1996) sought to further define the spectral sensitivity of TdR and Leu incorporation in natural bacterial populations. Incubations with TdR and Leu of surface waters from a mesotrophic estuary were performed on six separate occasions. Following exposure for 6.5 to 9 h, significant differences in inhibition of incorporation of TdR and Leu relative to a dark control for treatments in full sunlight and with UVB excluded occurred. Full sunlight and UVB inhibited TdR incorporation by 44%, and 39(7,, respectively. In contrast, Leu incorporation was inhibited 29% by full sunlight but 83(,;~ of this inhibition was due to UVB. Since bacterial production is inhibited significantly by UVB, it is likely that a major cause of this inhibition is direct damage to DNA. It has been shown that bacterioplankton may experience significant amounts of DNA damage (CPDs) in surface waters, often twice the amount of larger eukaryotic cells (Jeffrey et al., 1996a). Bacterioplankton have been shown to accumulate DNA damage over a solar day and to repair the majority of that damage during the night in the Gulf of Mexico (Jeffrey et al., 1996a,b). DNA damage may extend to depths of 10 in or more in calm waters but the amount of damage may be significantly altered by surface water mixing events (Jeffrey et al., 1996a; 1997). Radioimmunoassay (RIA) is a competitive binding assay between an unlabeled and radiolabeled antigen for binding to an antibody raised against that antigen. We have adapted this technique to the measurement of specific DNA photoproducts in the DNA of UV-irradiated cells (Mitchell and Clarkson, 1981; Mitchell, 1996). The following description is given for quantification of CPDs and (6-4) photoproducts in DNA using RIA. For convenience, the radiolabeled antigen is referred to as the 'probe' and the unlabeled competitor as 'sample' or 'standard'. The arnount of radiolabeled antigen bound to the antibody is determined by separating the antigen-antibody complex from free antigen by, for example, secondary antibody or high salt precipitation. The amount of radioactivity in the antigen-antibody complex in the presence of known amounts of competitor (i.e. standards) can then be used to quantify the amount of unknown sample present in the reaction. Under these conditions, 471
antibody binding to an unlabeled competitor (sample DNA) results in reduced binding to the radiolabeled ligand (i.e. inhibition). Samples are compared to results obtained with standard DNA (i.e. pUC19 plasmid DNA) irradiated (i.e. using UVC light) such that the frequency of photoproduct formation is known. DNA damage results are reported per unit (megabase) DNA and are therefore independent of the concentration of DNA present in the original sample or the amount of DNA assayed. The sensitivity of the RIA is determined by the affinity of the antibody and specific activity of the radiolabeled antigen (probe). Using high affinity antibody and probe labeled to a high specific activity, the reaction can be limited to such an extent that extremely low levels of damage in sample DNA can be detected.
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PRINCIPLE AND METHODOLOGY Commercial sources available for the purchase of anti-UV DNA antisera include Kamiya Biomedical Co., Thousand Oaks, CA (CPD monoclonal antibody) and Dermigen, Inc., Smithville, TX [CPD, (6-4) photoproduct, and custom rabbit antisera]. The following is a procedure for producing polyclonal antisera in rabbits. The following factors determine the affinity of polyclonal antisera in rabbits: (1) the 'foreigness' of the immunogen to the host (e.g. steric and distortive deviations from normal DNA structure); (2) the number of antigenic determinants presented to the host immune system (i.e. concentration, dose); (3) accessibility of the host immune system to the antigen (e.g. single or double strandedness of the DNA); (4) stability of the immunogen in the host animal; and (5) host variability. UVC radiation (240-290nm) produces multiple types of dimeric damage in DNA, predominantly the CPD, (6-4) photoproduct, and Dewar pyrimidinone (Cadet and Vigny, 1990). In a mixture of antigenic determinants, the lesion with the greatest immunogenicity will elicit the greatest immune response. The (6-4) photoproduct, which bends the DNA helix approximately 42 ° elicits a much greater response in rabbits than the CPD which bends DNA only about 7 °. The anti-(6-4)photoproduct subpopulation displays about 10- to 100-fold more affinity than the CPD. Because of this, UVC-DNA antisera can be diluted to such an extent that binding to the minor CPD antibody subpopulation is undetectable and the RIA is specific for the (6-4) photoproduct. For CPD antisera it is necessary to produce DNA containing this photoproduct exclusively. DNA is irradiated with UVB light (290-320 nm) in the presence of a triplet sensitizer (e.g. 2 x 10 ~ M acetophenone or 10~ acetone) to produce predominately cis,syn CPDs (Lamola and Yamane, 1967). The DNA is extensively dialyzed post-irradiation to remove the sensitizer. For production of anti-UV DNA antisera we typically immunize four rabbits and select the animals that show the greatest immune response for exsanguination (Figure 23.2). 472
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Equipment and reagents • 1-120 (HPLC or Millipore-filtered) • Salmon testes (or calf thymus) DNA (Sigma) • Acetone • Methylated bovine serum albumin (Sigma A1009) • DNA nick-translation kit (Boehringer-Mannheim #976 776) • 32P-labeled deoxynucleotide triphosphates (dNTPs) (NEN, Amersham, or
ICN) • TE buffer (I 0 mM Tris, pH 8.0, I mM EDTA) • Lysis buffer B [10 mM -Iris, pH 8.0, I mM EDTA, 0.5% SDS, 100 IJg ml' DNase-free RNase A (Boehringer-Mannheim #109-169)]
473
• Chloroform:isoamyl alcohol (24:1)
• Tris-saturated phenol (pH > 7.6) (Boehringer-Mannheim #100-997) • 10×TES (100 mMTris, pH 8.0, 10 mM EDTA, 1.5 M NaCI) • Gelatin (Type B: bovine skin) (Sigma #G-9382) • RIA buffer ( I x T E + 0.2% gelatin)The gelatin is heated into solution using a heated magnetic stirrer (not microwave) and heated to precisely 39J,0°C. Overheating (by as much as I-2°C) will result in prohibitive background! The cause of this is unknown. • Normal rabbit sera (NRS) (Calbiochem #566442); stored frozen in 200 IA aliquots.We have found that NRS (Calbiochem) diluted 1/40 in RIA buffer is optimal for immune pellet formation. Obviously other sources are readily available, however, we suggest that each batch be titrated in a binding assay to determine the optimal dilution. • Goat anti-rabbit IgG [Calbiochem #539844 or #539845 (bull<)]; stored frozen in 0.5 ml aliquots. • Tissue solubilizer (NCS-II from Amersham; #NNCS.502) supplemented with 10% (v:v) H20. • Scintillation cocktail (e.g. ScintiSafe from Fisher) containing I ml I' acetic acid (to neutralize the tissue solubilizer).
Preparation of i m m u n o g e n C o m m e r c i a l D N A (salmon testes or calf t h y m u s from Sigma) is diluted to lmgml' in 10ml sterile H,O (as d e t e r m i n e d by optical density at 260 nm). Diluted d o u b l e - s t r a n d e d D N A is UV-irradiated using one of the following protocols. 1. The i m m u n o g e n for anti-CPD sera is p r o d u c e d by irradiating the D N A solution (1 m g ml ') diluted in 10% acetone (final concentration; v:v) with - 7 5 k J m ~ UVB light in a glass 100 m m plate. The UVB source consists of four Westinghouse FS20 s u n l a m p s filtered through cellulose acetate (Kodacel f r o m Kodak) with a w a v e l e n g t h cutoff of 290 n m (Rosenstein, 1984). D o s i m e t r y is d e t e r m i n e d with an appropriate p h o t o m e t e r / r a d i o m e t e r (e.g. ILl400 p h o t o m e t e r coupled to a SCS 280 probe). At a distance of - 10 cm the fluence rate is - 5 J m -~s ~, hence, exposure times of N 4 h are required for a d e q u a t e CPD induction. The D N A is extensively dialyzed post-irradiation to r e m o v e a n y acetone. 2. The i m m u n o g e n for anti-(6-4)PD sera is p r o d u c e d b y irradiating D N A with 60 kJ m 2 UVC light. The UVC source consists of a b a n k of five Philips Sterilamp G8T5 bulbs emitting p r e d o m i n a n t l y 254 n m light. At a distance of ~ 20 cm the fluence rate is - 14 J m ~s ' and at this fluence rate the a v e r a g e duration of exposure is - 2 h. H e a t d e n a t u r e UV-irradiated D N A at 100°C for 10 min then place on ice. Single-stranded UV-irradiated D N A is then electrostatically coupled to m e t h y l a t e d bovine s e r u m albumin. Methylated BSA is a d d e d d r o p w i s e (approximately 50 pl each drop) until the UV-irradiated D N A solution turns cloudy.
474
Immunization schedule Four New Zealand White female rabbits are initially injected subcutaneously at 10 sites (100 pl each) with 0.5 ml immunogen mixed with an equal volume of Freund's Complete Adjuvant (final concentration of 0. I mg ml ' UV-DNA). Rabbits are subsequently injected using the same protocol as above at two week intervals except that Freund's Incomplete, rather than Complete, Adjuvant is mixed with 0.5 ml immunogen. At 10-12 days following the second injection I ml of serum is drawn and binding affinity evaluated using immunoprecipitation (see below). Immunization is continued at two week intervals until sufficient binding activity is attained at which time antisera (60-80 ml) are drawn from the animal using heart puncture. Antisera are dispensed into I ml aliquots and stored at -20°C. Repeated freezing and thawing of antisera is to be strictly limited since this can severely reduce binding activity.
Determination of antiserum binding using immunoprecipitation • Probe synthesis: Both CPD and (6-4)PD frequencies are greatest in nucleic acid substrates containing a high A+T:G+C ratio. Hence, optimal substrates for the radiolabeled probe include Clostridium perfringens D N A as well as the homopolymer poly(dA):poly(dT). Nick-translate D N A (0. I lag) with ~2P-dCTP and/or 3~P-TTP to give a specific activity of - 5 × 108-109cpm lag ' (BoehringerMannheim NickTranslation Kit #976 776).A typical reaction includes 2 pl 10× buffer (from kit); 2 lal dATP [for poly(dA):poly(dT)]; or 2 tJI each dATP and dGTP (for DNA); 0.5 lal poly(dA):poly(dT) or C. perfringens D N A (diluted to 20 lug per 100 lal); 12.5 la132P-TTP at 10 mCi ml ' (NEN orAmersham); 3 4 lal DNasel/DNA polymerase I enzyme mix (from kit). Incubate for 3 0 4 5 min at 15°C. Separate radiolabeled ligand from free dNTPs using a Nick Column (Pharmacia) eluting the ligand with TE buffer. • Irradiate 32P-labeled probe with 30 kJ m 2 UVC light and restore the volume if necessary (due to evaporation) with H20 and dilute 2500- to 5000-fold in RIA buffer (yielding 2.5-5.0 pg probe in 50 pl buffer).The amount of probe added to the RIA determines its sensitivity. It is essential to use 10 pg or less and have enough radioactivity in the assay (cpm) to yield useful binding (and inhibition) data. Hence, if a 5000 dilution of probe leaves < 500 cpm in 50 lal, a greater concentration must be used. Good assay conditions should be limited to at least 500 cpm per 50 lal (added to reaction) at a probe dilution not to exceed I/I 250. • Add I ml of RIA buffer to duplicate 12 mm disposable culture tubes. Add 50 lal antiserum diluted in RIA buffer at half-log increments from 1/1000 to I/I 000 000 (dilution prior to dispensing). Duplicate tubes without antiserum are dispensed to determine background. Add 50 pl of diluted ~2P-labeled probe and vortex well. Incubate for 3 4 h with gentle rotation (optional) in a 37°C dry incubator. • Separately add 50 lal of normal rabbit serum diluted 1/40 in RIA buffer and 50 lal of goat anti-rabbit IgG diluted 1/20 and vortex well. Incubate at 4°C for 2 days until immune pellet (translucence) develops. Centrifuge tubes at - 35004000 rpm for 30 to 45 rain. Decant supernatant, invert tubes onto absorbant paper in
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test tube rack, and drain for 5 min. Wipe the lip of the tube with a cotton applicator covered with a tissue to remove any accumulated liquid. Add 100 lul of NCS Tissue Solubilizer (Amersham) supplemented with 10% H20 and incubate at 37°C (or room temperature) with rotation until immune pellet is completely dissolved. It is extremely important that the immune pellet be completely solubilized but not allowed to dry. Partial solubilization will result in bad duplicates. Add 2 ml of scintillation cocktail (e.g. ScintiSafe from Fisher) supplemented with I ml I ' acetic acid (to eliminated chemoluminescence generated by tissue solubilizer) and vortex.The samples can either be decanted into 20 ml scintillation vials and washed twice with 4 ml additional scintillation cocktail or the RIA tubes can be placed directly into scintillation vials and counted. Count 32p using liquid scintillation counter.
Isolation of D N A One of the major attributes of RIA is its ability to measure photoproducts in DNA that has not been extensively purified and most DNA isolation protocols are suitable for RIA sample preparation. The protocol described below has been found to work well in a variety of marine environments. The radioimmunoassay is less dependent on DNA concentration than on the number of photolesions present. If DNA damage is low, more cells will be needed. Conversely, smaller samples may be used when damage is high. In general, we have found that a final extract of 2-4 pg DNA is sufficient to detect damage in marine planktonic communities, although 5-10 pg is preferable. This allows replicate analysis for each photoproduct as well as multiple photoproducts. In open ocean samples, this may require 50-701 of seawater. In coastal waters, 51 is often sufficient. In general, samples are collected by filtration. Size fraction filtration may be used to isolate particular members of the microbial community (e.g. bacterioplankton <0.8 l~m, >0.21.lm pore size). We have found Gelman SUPOR (polysulfone) filters with 142 mm diameters to work well. Filters are collected, folded, and placed in 2 oz polyethylene bags. Filters are stored at -80°C until extraction. To extract DNA remove the filter from the freezer and immediately crush the filter in the bag before it thaws. Filter pieces were then poured into a 50 ml Oak Ridge centrifuge tube. • Add 5 ml of STE (I 0 mM Tris, pH 8; I mM EDTA; I00 mM NaCI) containing 1% SDS (sodium dodecyl sulfate) to each tube. Cap, vortex for 15 s, and place in a boiling water bath for 5 rain. • Aspirate the lysate and place in a new centrifuge tube.Wash the filter pieces with an additional 5 ml of STE, vortex for 15 s, and combine with the lysate. • Extract with I 0 ml of chloroform:isoamyl alchohol (24:1). Collect the aqueous phase after centrifuging at 4°C for 30 min at 3000 rpm. • Decant the aqueous phase into a new Oak Ridge tube, add I 0 lal of a 25 mg ml ' glycogen solution, 0. I volume of 3 M sodium acetate (pH 5), and an equal volume of isopropanoI.Vortex and precipitate the sample overnight at -20°C. • Collect the precipitate by centrifugation at 12000g and 4°C for 30 rain. Decant the supernatant and wash the pellet with 10 ml cold 70% ethanol. Centrifuge at 12 O00g at 4°C for I 0 min. Decant the supernatant.
476
• Re-suspend the pellet in I ml of STE before transferring to a 2 ml microfuge tube. Re-precipitate as above, dry the pellet then re-suspend in 200 pl STE. DNA concentrations are determined spectrofluorometrically (Paul and Myers, 1982) using either a Picogreen (for native DNA) or Oligreen (for denatured DNA) reagent from Molecular Probes (Eugene, OR).We prefer the Oligreen method since it quantifies DNA concentrations immediately prior to analysis.
Competitive binding assay (RIA) The RIA is s i m p l y the basic i m m u n o p r e c i p i t a t i o n reaction outlined a b o v e into which a standard or s a m p l e D N A has been a d d e d to c o m p e t e with the radiolabeled p r o b e for a n t i b o d y binding. Hence, the p r o c e d u r e is exactly the s a m e as that used for i m m u n o p r e c i p i t a t i o n with the following additions/modifications. • A single dilution of antiserum is used. This dilution is determined from immunoprecipitation analyses of binding activity (see above) and should yield 30-60% of the radiolabeled probe in the immune pellet. • For quantification of CPDs or (6-4)PDs a dose response of heat-denatured salmon testes (Sigma) UV-irradiated DNA is used as standard (Table 23. I).We routinely use doses of 3, I 0, 30, 100, and 300 J m 2 as our standard curve and assay the same amount of standard as sample DNA.We have determined rates of photoproduct induction using independent analysis of CPDs as T4 UV endonuclease-sensitive sites and (6-4) photoproducts as photoinduced alkalilabile sites (Mitchell et al., 1990). From these values the UVC doses are converted to photoproduct frequencies using 8. I CPDs and 1.56 (6-4) photoproducts per megabase DNA per joule m 2.When relative, rather than exact, amounts of CPDs or (6-4)PDs are adequate for experimental purposes (as in DNA repair experiments) the sample harvested at the time of irradiation can be diluted in half-log increments to generate a standard. From this type of standard curve data representing percentage photoproducts remaining (or repaired) can be generated. • Unlabeled competitor mammalian DNA, radioactive ligand, and diluted antibody are incubated together for 3 h at 37°C with gentle rotation (optional). (As above, it is prudent to perform a preliminary titration of sample DNA to determine the amount required for adequate inhibition in the RIA).The total volume of sample DNA added can vary within certain limitations. Sample volumes <100 [JI do not significantly effect the reaction conditions (e.g. total binding). Sample volumes > 100 pl can be used, however, the total reaction volume should be increased accordingly (i.e. doubled).
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APPLICATIONS We h a v e used the described m e t h o d to s t u d y D N A d a m a g e in a wide variety of systems, from the Southern Ocean (Jeffrey et al., 1997) to coral reef microbial c o m m u n i t i e s (Lyons et al., 1998). Tlle protocol w o r k s
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equally well for bacterioplankton, phytoplankton (Karentz et al., 1991), and fish larvae (Malloy et al., 1997). Most often, the technique has been used to determine the amount of UVB-induced DNA damage in microbial communities (Jeffrey et al., 1996a,b). This has been done as a function of time of day, depth in the water column, cell size fraction, or seasonal comparisons. It is also possible to examine rates of DNA damage repair by incorporating broadband spectral filters (e.g. Mylar 500 D or acrylics, Aas et al., 1996) post-UVB irradiation. The main obstacle to doing this with marine microbial communities is the large volume required to be filtered for each time point. We have had some success by constructing UV-transparent incubators that will hold up to 200 1of seawater at ambient temperatures from which samples may be collected over time. High cell densities in laboratory cultures allow more detailed experimentation to be designed (Joux et al., 1999). The DNA damage assays are identical, with only minor modifications in the DNA extraction protocol required. The kinetics of DNA damage induction and repair may be determined as a function of dose response, for instance, and comparisons made between different strains. Smaller volumes may allow greater spectral resolution of a UV response by incorporating additional optical filters (e.g. Schott filters, Schott Glass Technologies, Duryea, PA) which are not available in sizes needed to incubate seawater samples.
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DATA ANALYSIS A sample Excel spreadsheet for quantification of CPDs or (6-4)PDs is shown in Table 23.1. A sample Excel spreadsheet for quantification of relative photoproducts remaining at specific times post UV-irradiation (e.g. in a DNA repair experiment) is shown in Table 23.2.
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CONCLUSIONS
AND FUTURE DIRECTIONS
Different types of DNA damage have different immunogenicities and may not be suitable for the production of antibodies in rabbits or mice. We have been unsuccessful in raising rabbit antisera against DNA irradiated with UVA light, DNA cross-linked with mitomycin C, or DNA homopolymers or alternating copolymers irradiated with UVC light. These failures may be due to the low antigenicity of the damage itself (e.g. UVA, mitomycin C), immunogen stability in the host (e.g. digestion of oligonucleotides or polynucleotides), or host variability (i.e. not enough rabbits). We have been successful, however, at raising polyclonal antibodies against UVC light, triplet-sensitized UVB light, DNA containing acetylaminofluorene adducts, DNA containing benzo[a]pyrene diolepoxide adducts, DNA treated with osmium tetroxide (i.e. thymine glycols), and HPLC-purified 8-oxodeoxyguanosine covalently linked to a protein hapten. 480
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RIA and ELISA are p o w e r f u l techniques for the sensitive and specific detection of genotoxic d a m a g e in DNA. Other techniques, such as quantitative i m m u n o c y t o c h e m i s t r y , immunohistochemistry, and i m m u n o e l e c tron m i c r o s c o p y have been designed to detect D N A d a m a g e in situ, thus visualizing the distribution of d a m a g e in tissues and cells. D a m a g e d D N A m a y be separated from u n d a m a g e d D N A fragments b y i m m u n o precipitation, in-tmunoaffinity chromatography, or nitrocellulose binding. Enrichment of d a m a g e d D N A f r a g m e n t s using antibodies has been c o m b i n e d with PCR amplification and Southern analysis to determine the genomic distribution of D N A d a m a g e (Hochleitner et al., 1991) and with HPLC (or ligation-mediated PCR) to increase the resolution and, hence, sensitivity of detection ( G r o o p m a n et al., 1992).
References Aas, P., Lyons, M., Pledger, R., Mitchell, D. L. and Jeffrey, W. H. (1996). Inhibition of bacterial activities by solar radiation in nearshore waters and the Gulf of Mexico. Aquatic Microbial Ecol. 11, 229-238. Cadet, J. and Vigny, P. (1990). The photochemistry of nucleic acids. In: Biooor~aJTic PhotochenlistJ 7, Vol. 1: Ptlotochenlistry and the Nuch'ic Acids (H. Morrison, Ed.), pp. 1-272. John Wiley and Sons, New Yurk. Calkins, J. (Ed.) (1982). The Roh' qf Solar Llltravi~det RadiatioH iH Marine Ecosystems. Plenum Press, New York. Chin-Leo, G. and Kirchman, D. L. (1988). Estimating bacterial production in marine waters from the simultaneous incorporation of thymidine and leucine. Appl. Euviron. Micmbiol. 54, 1934 1939. Coffin, R. B., Velinsky, D., Devereux, R., Price, W. A. and Cifuentes, L. (1990). Stable carbon isotope analysis of nucleic acids to trace sources of dissolved substrates used by estuarine bacteria. Appl. Euvirol~. Micmbiol. 56, 2012-2020. Cota, G. E, Kottmeier, S. T., Robinson, D. H., Smith, W. O. and Sullivan, C. W. (1990). Bacterioplankton in the marginal ice zone of the Weddel] Sea: biomass, production and metabolic activities during Austral summer. Deep-Sea Res. 37, 1145-1167. Falkowski, P. G. and Woodham, A. D. (1992). Prima12t/Production aud Biogeochemical Cycles in the Sea. Plenum Press, New York. Fuhrman, J. A. and Azam, E (1980). Bacterioplankton secondary production estimates for coastal waters of British Columbia, Antarctica, and California. App[. El~virou. Microbiol. 39, 1085 1095. Fuhrman, J. A. and Azam, E (1982). Thymidine incorporation as a measure of heterotrophic bacterioplankton production in marine surface waters: evaluation and field results. Mar. Biol. 66, 109-120. Groopman, J. D., Zhu, J. Q., Donahue, P. R., Pikul, A., Zhang, L. S., Chen, J. S. and Wogan, G.N. (1992). Molecular dosimetry of urinary aflatoxin-DNA adducts in people living in Guangxi Autonomous Region, People's Republic of China. Ca~Tcer Res. 52, 45-52. Hansen, R. B., Sharer, D., Ryan, T., Pope, D. and Lowery, H. K. (1983). Bacterioplankton in the Antarctic ocean waters during late Austral winter, abundance, frequency of dividing cells, and estimates of production. Appl. EllviroJl. Microbiol. 45, 1622 1632. Herndl, G. ]., M/_iller-Niklas, G. and Frick, J. (1993). Major role of ultraviolet-B in controlling bacterioplankton growth in the surface layer of the ocean. Nature 361, 717-719.
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Hochleitner, K., Thomale, J., Nikitin, A. Yu and Rajewsky, M. E (1991). Monoclonal antibody-based, selective isolation of DNA fragments containing an alkylated base to be quantified in defined gene sequences. Nucleic Acids Rcs. 19, 4467-4472. Jeffrey, W. H., Aas, P, Lyons, M. M., Pledger, R., Mitchell, D. L. and Coffin, R. B. (1996a). Ambient solar radiation induced photodamage in marine bacterioplankton. Photochenl. Photobiol. 64, 419-427. Jeffrey, W. 1t., Miller, R. V. and Mitchell, D. L. (1997). Detection of ultraviolet radiation induced DNA damage in microbial communities of the Gerlache Strait. Aittarctic J. LIS 32, 85-87. Jeffre}; W. H., Pledger, R. J., Aas, P., Hager, S., Coffin, R. B., Von Haven, R. and Mitchell, D. L. (1996b). Diel and depth profiles of DNA photodamage in bacteriuplankton exposed to ambient solar radiation. Mar'. Ecol. Prog. Ser. 137, 283-291. Joux, E, Jeffrey, W. H., Lebaron, I~ and Mitchell, D. (1999). Marine bacteria display diverse responses to u/traviolet-B radiation. Appl. Environ7. Microbiol. 65, 3820-3827. Karentz, D., Cleaver, J. E. and Mitchell, D. L. (1991). Cell survival characteristics and molecular responses of antarctic phytoplankton to ultraviolet-B radiation. J. Phycol. 27, 326-341. Lamola, A. A. and Yamane, T. (1967). Sensitized photodimerization of thymine in DNA. Proc. Natl. AcmL Sci. LISA 58, 443-446. Lyons, M. M., Aas, P., Pakulski, J. D., Van Waasbergen, L., Mitchell, D. L., Miller, R. V. and Jeffrey, W. H. (1998). Ultraviolet radiation induced DNA damage in coral reef microbial communities. Mar. Biol. 130, 537-543. Malloy, K. D., Holman, M. A., Mitchell, D. and Detrich, H. W. III (1997). Solar UVBinduced DNA damage and photoenzymatic repair in Antarctic zooplankton. Proc. N~ttl. Acad. Sci. USA 94, 1258-1263. Mitchell, D. L. (1995). Ultraviolet radiation damage to DNA. In: Molecular Biology amt Bioteclmology: A Comptvhensive Desk Reference (R. A. Meyers, Ed.), pp. 939-943. VCH Publishers, New York. Mitchell, D. L. (1996). Radioimmunoassay of DNA damaged by ultraviolet light. In: Techllologics (or Detectiol~ qf DNA Damage ~TmtMutatiotzs (G. Pfeifer, Ed.), pp. 73-85. Plenum, New York, Mitchell, D. L. and Clarkson, J. M. (1981). The development of a radioimmunoassay for the detection of photoproducts in mammalian cell DNA. Biochim. Biophys. Acta 655, 54 60. Mitchell, D. L. and Nairn, R. S. (1989). The biology of the (6-4) photoproduct. Photochem. Photobiol. 49, 805-819. Mitchell, D. L., Brash, D. E. and Nairn, R. S. (1990). Rapid repair of pyrimidine (6-4)pyrimidone photoproducts in human cells does not result from change in epitope conformation. NHcleic Acids Res. 18, 963-971. Paul, J. H. and Carlson, D. (1984). Genetic material in the marine environment: implication for bacterial DNA. Limm~l. Oceam~gr. 29, 1091-1097. Paul, J. H. and Myers, B. (1982). Fluorometric determination of DNA in aquatic microorganisms by use of Hoechst 33258. Appl. E1~virott. Microbiol. 43, 1393-1399. Paul, J. H., Jeffrey, W. H. and Deflaun, M. E (1985). Particulate DNA in subtropical oceanic and estuarine planktonic environments. Mar. Biol. 90, 95-101; Appl. EHvirotz, Microbio[. 43, 1393-1399. Rosenstein, B. S. (1984). Photoreactivation of ICR 2A frog cells exposed to solar UV wavelengths. Photochcm. Photobiol. 40, 207-213. Simon, M. and Azam, E (1989). Protein content and protein synthesis rates of planktonic marine bacteria. Mar. Ecol. Pro~,,. Set. 51,201-213.
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Z
!d i
Sullivan, C. W., Cota, G. E, Krempin, D. W. and Smith, W. O. Jr. (1990). Distribution and activity of bacterioplankton in the marginal ice zone of the Weddell-Scotia Sea during austral spring. Mar. Ecol. Prog. Set. 63, 239-252. Vincent, W. E and Roy, S. (1993). Solar ultraviolet-B radiation and aquatic primary production: damage, protection, and recovery. Environ. Rev. 1, 1-12. Worrest, R., Dyke, H. V. and Thomson, B. (1978). Impact of enhanced simulated solar ultraviolet radiation upon a marine community. Photochem. Photobiol. 27, 471-478. Worrest, R., Wolniakowski, K., Scott, J., Brooker, D. and Dyke, H. V. (1981). Sensitivity of marine phytoplankton to UV-B radiation: impact upon a model ecosystem. Photochem. Photobiol. 33, 223-227. Zepp, R., Callaghan, T. and Erickson, D. (1995). Effects of increased solar ultraviolet radiation on biogeochemical cycles. Ambio 24, 181-187.
List of suppliers Amersham Pharmacia Biotech, Inc. 800 Centennial Avemte P.O. Box 1327 Piscataway, NJ 08855-1327, USA Tel: 732-457-8000 Fax: 732-457-0557 Radio-labeled dNTPs; Tissue solubilizer
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Liquid scintillation cocktail
Calbiochem-Novabiochem Corporation 10394 Pacific Center Court San Diego, California 92121, USA Mailing Address: P.O. Box 12087 La Jolla, California 92039-2087, USA Teh 858- 450-9600 Fax: 858- 453-3552 N o r m a l rabbit sera; Goat antirabbit (IgG)
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Sigma Chemical 3050 Spruce Street St. Louis, MO 63103, USA Tel: 314-771-5765
DNA; methylated BSA; gelatin
Oligogreen nucleic acid dye New England Nuclear (NEN) NEN® Life Science Products, Inc. 549 Albany Street Boston, MA 02118-2512, USA Teh 800-551-2121 or 617-482-9595 Fax: 617-482-1380
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485
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24 Radiotracer Assays of Sulfate Reduction Rates in Marine and Freshwater Sediments G a r y M King Darling Marine Center,University of Maine, Walpole, ME 045 73, USA
CONTENTS Introduction Brief history of ~SO~~ methodologies Single-step chromium reduction for assay of sulfate reduction rates Modifications of the single-step procedure
INTRODUCTION Sulfate dominates the dissolved electron acceptors in marine sediments, with concentrations (up to 28 raM) much greater than those of all others (e.g. oxygen, nitrate) combined. Freshwater sulfate concentrations are also relatively high, often exceeding 100 btM. In contrast, oxygen and nitrate concentrations seldom exceed 300 ~M and 100 btM, respectively, in freshwaters. Thus, once oxygen is depleted, dissimilatory sulfate reduction dominates anaerobic metabolism in marine systems (predominantly in sediments) and often plays an important role in freshwater systems (Capone and Kiene, 1988) and algal mats (Frtind and Coben, 1992). In addition, transformations of sulfate reduction endproducts (e.g. dissolved and metal sulfides) play an important role in oxygen budgets and can limit oxygen availability for aerobic heterotrophs (Jorgensen, 1982; Fenchel et al., 1998). Much of the current understanding of sulfate reduction has been derived from assays based on ~SO4e , a readily available and relatively inexpensive radiotracer that can be purchased in a carrier-free form with very high specific activity. However, prior to routine use of radiolabeled sulfate, sulfate reduction rates in marine sediments were assayed successfully with depletion methods and diagenetic modeling (Berner, 1964; Martens and Berner, 1977; Jorgensen, 1978a,b; Canfield, 1989). Depletion approaches, which simply measure loss of sulfate over time, have proven M E T H O D S IN MICROBIOLOGY, VOLUME 30 1SBN 0 12 521530-4
C o p y r i g h t © 2001 Academic Press Ltd All rights of reproduction in any form reserved
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suitable in cases where extended incubations (days to weeks) are feasible. Furthermore, depletion approaches, which have no significant regulatory concerns (e.g. disposal of radioactive wastes) and which require minimal sample processing and analytical facilities, have offered attractive nonisotopic alternatives to radioassays. Diagenetic modeling, which is based on the use of diffusion-reaction models and sulfate and organic matter concentrations, has been particularly useful for estimating sulfate reduction in deep-sea sediments, where activity occurs at low rates over many meters of the sediment column (Berner, 1964). Diagenetic modeling has also been applied effectively in coastal systems where concentration gradients occur over intervals of tens of centimeters (Jorgensen, 1978a,b). Though an alternative to isotopic assays, diagenetic modeling assumes steady-state conditions that often do not obtain in coastal systems. In addition, diagenetic models often require information on or assumptions about sedimentation rates, diffusion coefficients, bioturbation and other ancillary parameters. In spite of the inherent limitations on the use of any radiotracer, sulfate reduction assays using ~;SO~e may be preferable for many marine and freshwater systems, including microbial mats. Advantages of ;~SO~-' include a high level of sensitivity, even for very low rates of activity, the possibility of assessing activity in the presence of sulfide oxidation, short incubation times under near in situ conditions and information on endproduct formation (e.g. elemental sulfur, pyrite). Relatively simple modifications of the methodology for '~SO,2 also provide options for assessing transformations of other sulfur species (e.g. S", thiosulfate, metal sulfides).
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BRIEF H I S T O R Y O F
355042- M E T H O D O L O G I E S
Ivanov (1956) first used ~SO~e to determine sulfate reduction rates in various marine sediments and developed the basic methodology that has been adopted for contemporary studies. Briefly, mixed sediments or water column samples are incubated anaerobically after addition of 3~SO~2 stocks. H2~Sresulting from sulfate reduction is distilled after acidification and trapped for subsequent radioassay. Jorgensen, Skyring and others (see Jorgensen, 1978a,b; Skyring, 1987 and references therein) have modified this method and popularized 'direct injection' techniques that introduce 3~SO42 as a true tracer (i.e. in a carrier-free form) into largely undisturbed sediments. Cohen (personal communication) has described a methodological variant in which silver wires are used as a vehicle for introducing radiosulfate and trapping radiosulfide. This approach can provide rate information, but is also a useful qualitative tool for mapping the distribution of sulfate reduction on a fine scale (millimeter). Carrier-free ~SO;2 stocks facilitate assays with undisturbed samples since it can be reasonably assumed that the fraction of ~SO4~ reduced anywhere within a sample reflects the local rate of sulfate reduction independently of small-scale variations in sulfate concentrations and the 490
amount of isotope present. As a result, an integrated sulfate reduction rate for a sample can be obtained by evaluating the following expression or a variant thereof: sulfate reduction rate = ([H~%/%O~ ]*~-SO~ )*IDF/T where concentrations in units suitable for a given sample (e.g. dpm ~S cm ~sediment or nmol '~SO42 cm ; sediment) are indicated in brackets, T = incubation time and IDF = an isotopic discrimination factor, sometimes ignored and otherwise taken as 1.06. For many earlier studies, HCl-based distillations had been considered adequate for volatilizing dissolved radiosulfides and Fe'sS present in amorphous to moderately well-crystallized metal sulfide phases. However, several lines of evidence suggested that a variable portion of the ';SO4: reduced within a given sample occurs in forms not subject to acid distillation (Howarth and Teal, 1979; King, 1983; Howarth and Merkel, 1984; Howes el al., 1984; King et al., 1985). In particular, neither ~@' nor Fe-;%, can be distilled from a sample matrix by HCI alone. However, both of these species can be produced as artefacts of isotopic exchange, or in the case of "S~', by HC1 distillation per se (Fossing et al., 1992). While isotopic exchange can vary from negligible to significant levels depending on several factors, HC1 distillation invariably releases soluble ferric iron from many sediments, resulting in a rapid chemical oxidation of H2~% to ~S". In addition, both '~S" and Fe~S~ can be formed as a consequence of processes in situ, on occasion accounting for a substantial fraction of reduced radiosulfur. Obviously, accurate sulfate reduction rate estimates require analysis of radioactivity in all relevant sulfur pools. Several options have been used to assess radioactivity in non-acid volatile reduced sulfur fractions. ~S*' has been measured specifically by extracting elemental sulfur with solvents such as hexane, benzene or carbon disulfide (Troelsen and ]orgensen, 1982; King, 1988). The latter is the most effective extractant, but also the most hazardous with respect to health and flammability and should be used with considerable caution. Subsequent to acid distillation to measure volatile sulfides and extensive washing to remove unreacted radiosulfate, aqua regia (a 1:1 mixture of concentrated hydrochloric and nitric acids) digestions have been used to solubilize both elemental sulfur and pyrite (as well as organic sulfur) in sediments. However, aqua regia digestions have proven somewhat controversial, yielding inconsistent results (Howarth and Giblin, 1983; King, 1983; Howarth and Merkel, 1984; Howes ct al., 1984; King et al., 1985). In addition, aqua regia digestions are cumbersome, hazardous and generate substantial amounts of radioactive waste. As an improvement, boiling acidic chromous chloride distillations have proven reliable and more practical. The basic methodology was introduced by Zhabina and Volkov (1978) and later Westrich (1983) to fractionate various inorganic reduced sedimentary sulfur pools. The key feature of the method is that acidic chromous chloride reduces inorganic sulfur species, including pyrite but excluding sulfate, to hydrogen sulfide. Although the method has been used with sequential HC1 distillation and 491
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carbon disulfide extraction to assay acid volatile sulfur, elemental sulfur and pyritic sulfur fractions, respectively (Howes et al., 1984; King et al., 1985; King, 1988), single-step processing has become routine since quantitative estimates of elemental and pyritic sulfur formation rates are often precluded by isotopic exchange and other processing artefacts (as above). Thus, single-step chromous chloride distillation is emphasized in the following method description (see also King, 1988; Fossing and Jorgensen, 1989; King, 1990).
SINGLE STEP CHROMIUM REDUCTION FOR A S S A Y O F S U L F A T E R E D U C T I O N RATES Principle: All forms of reduced inorganic sulfur (i.e. non-sulfur), including pyritic sulfur and elemental sulfur, can be reduced by a solution of acidic chromous chloride to hydrogen sulfide. The latter is distilled from sediment or suitable aqueous samples, and trapped for mass assay, radioassay or both.
Introduction of 35SO42-into samples, incubation and termination Several options exist for collecting samples and adding 3~SO4~ to them. A common approach for sediments involves the direct injection of ~SO~: into sediments contained within acrylic core tubes with an inner diameter of about 2-3 cm (Jorgensen, 1978a,b; King and Klug, 1982). Intact, minireally disturbed samples can be collected by hand from shallow sediments, by SCUBA from deeper sediments, or as sub-cores from 'box cores' used when SCUBA is not feasible. The core tubes typically contain 1-2 mm ports drilled along a vertical axis at 1-2 cm intervals; prior to sampling, ports are sealed with silicone cement. Often, the sediment surface within a tube can be adjusted so that it extends a known distance above a port (0.5-1 cm), thus facilitating measurement of sulfate reduction rates with depth. A small volume (10 pl) of a solution of carrier-free ~SO4~ (370-3700 kBq ml ~ in deionized water or sterile seawater) is injected using a 10 ~1 syringe through ports at desired depth intervals. With practice, the injection can be made while withdrawing the syringe from the port, thereby creating a 'line' of radiolabel at a specific depth in the core. Injection times are noted and cores subsequently incubated at a suitable temperature for several hours (long incubations should be avoided). Intact cores from oxic systems can be incubated with a small layer of unstirred water (1-2 ram) above the sediment surface and an air headspace; cores from anoxic systems should be incubated sealed with a nitrogen headspace. Freezing cores rapidly terminates incubations. Access to an ultracold freezer (< -70°C) is preferable, but cores can also be placed in a -20°C freezer. In either case, a set of 'blank' cores should be frozen immediately after injection of ~'SO4: so that activity during the freezing stage can be assessed. Once frozen, cores can be held for brief periods (days) prior to 492
further processing. Frozen cores can be extruded to initiate processing; the desired intervals are sliced with a fine saw blade. These intervals are then treated as below. As an alternative, cores can be sectioned while fresh and the sections rapidly mixed into a solution of zinc acetate (~>5%) in a container suitable for sample storage. The zinc acetate rapidly terminates microbial activity and preserves sulfides. The resulting slurry is amenable to analysis as outlined below. In some instances, the coring approach described above m a y be impractical. In such cases, 'mini-cores' prepared by removing the luer tips from plastic disposable syringes m a y prove more suitable. These small (3-5 cm ~) piston cores can be used to obtain largely undisturbed samples at discrete depths from m u c h larger cores, collect intact sediments perpendicular to animal burrows, sample a limited vertical interval from cores or sediments in situ, or sample sediment slurries (King et al., 1990; Hansen et al., 1996; Giray and King, 1997). The syringe plunger serves as an effective seal for one end of the mini-core and a butyl septum can be used to seal the other. The septum also serves as a port for injecting ~SO, -~ along the axis of the mini-core. Mini-cores can be frozen quickly and easily in a dry ice-ethanol bath, are stored conveniently and can be expressed with little difficulty into boiling flasks for c h r o m i u m distillation (as below). They are also inexpensive, easy to prepare, help to minimize contamination, and can be discarded in solid radioactive waste; in contrast, acrylic core tubes are s o m e w h a t more cumbersome, p e r m a n e n t and require more care once contaminated with isotope.
Distillation, trapping and assay of reduced 3sS endproducts from sulfate reduction Essential hardware (see Figure 24.1). Reduction of non-sulfate ~S-sulfur and distillation of ~S-sulfides are best carried out in r o u n d - b o t t o m boiling flasks heated in sand baths or with heating mantles. The flasks are connected to a condenser through which cooling water flows at a moderate rate. The condenser outlet is constructed of a length of Teflon tubing (0.32 cm outer diameter or as appropriate) that connects a series of traps containing a solution of 5'Z( zinc acetate (7 ml per trap). Standard glass or plastic scintillation vials offer a convenient option for trap hardware since they are relatively inexpensive, disposable and facilitate radioassay with m i n i m u m additional handling. Several options exist for the design of an apparatus for distillation. Figure 24.1 illustrates one such design, in which three scintillation vials per boiling flask are connected in series. The vials are secured to an acrylic plate (1 cm thick or greater) into which long-skirt caps matching the thread of the vials have been secured with epoxy glue. These caps are each fitted with a Teflon-lined septum (20 m m diameter, Supelco, Inc.). Caps, septa and the acrylic plate are drilled to accept Teflon tubing, a length of which is used to connect the outlet of each condenser to a series of traps. The condenser outlet and tubing can be connected using a standard ground glass 'tubing connector'. Three-neck boiling flasks offer the greatest flexibility for attaching
493
roD
4,J
n-
H
F>
°
K
Figure 24.1. Schematic of a distillation apparatus for trapping chromium reducible sulfur. T-connector (A) leading from condenser (B) to a length of Teflon tubing (H); inlet and outlet for condenser cooling water indicated by G (note: several condensers may be connected in series). 100 or 200 ml three-neck boiling flask (C) with closure (D) on one port for adding sample and chromous chloride. Inlet (F) for a stream of nitrogen from a valve and flow regulator. Heating mantle or sand bath (E) for bringing chromous chloride to boiling. Scintillation vial (I) with zinc acetate mounted in an acrylic board (J) with vial caps recessed into the board. Threaded rod (K) mounted onto acrylic base or plywood base coated with polyurethane to facilitate cleaning.
condensers, gas inlets and s a m p l e ports. Flasks of 100ml or 250ml v o l u m e with 14/20 g r o u n d glass fittings are suitable for typical applications, d e p e n d i n g on sediment and c h r o m o u s chloride volume. Gas inlets can be p r e p a r e d using standard fittings that allow r e a d y connection to a manifold that is used to control a flow of nitrogen to each boiling flask. Flows are best regulated individually for each flask. On a three-neck boiling flask, the u n u s e d port is sealed with a ground glass p l u g that can be o p e n e d to introduce samples and c h r o m o u s chloride, following the p r o c e d u r e below.
Procedure 1. Assemble distillation glassware, establish a flow of cooling water, and prepare and position scintillation vial traps with a solution of zinc acetate (5-10%, 7 ml each). One or two d r o p s of Antifoam B or equivalent (Sigma Chemical Co.) should be a d d e d to the traps to prevent foaming. 2. Establish a flow of nitrogen (30-50 ml min ' for 10-15 min to pre-flush the system. 3. Transfer into r o u n d - b o t t o m boiling flasks containing 5 ml of 5% zinc acetate u p to 5 cm ' of a suitably p r e p a r e d s e d i m e n t s a m p l e with radiolabeled sulfides; s e d i m e n t can be a d d e d via a side-port as described above. Swirl gently to mix (this can be accomplished by loosening any fittings used to secure the condenser and flask assemblies).
494
4. Open side-port fittings while gassing with nitrogen and add chromous chloride to each boiling flask (see below) at 10 ml cm ~sediment. Apply heat and boil for 60 rain. Note: when adding chromous chloride, care must be taken that a partial
vacuum does not result from adding a cool solution to glassware warmed by a sand bath above ambient temperature; a vacuum can result in a flouT of zinc acetate from the scintillation vial traps into the boiling flasks. 5. Reduce heat after terminating the boiling step and remove the scintillation vial traps while maintaining a flow of nitrogen; this is best done by starting with the last trap in the series. 6. Sub-sample the scintillation vials as necessary for assays of total sulfide content [following Cline (1969) as described by King (1988) and othersl, and then add scintillation cocktail for radioassay [at least 1 : 1 by volume for Scintiverse II or 1 : 2 for Scintiverse BD (Fisher Scientific, Inc.)] using routine scintillation counting procedures and methods of standardization. Sum the radiolabel in the traps of a give series to estimate to distilled ~S. Alternatively, the zinc acetate solutions for each of the traps in a given series may be pooled and then sub-sampled for radioassay. The advantage of this approach is that small volumes of solution can be assayed in some cases with greater scintillation counting efficiencies; in addition, the approach facilitates use of 'mini-vials' for radioassay and smaller volumes of scintillation fluid. The disadvantages of the approach are that it requires significantly greater handling of radioactive solutions and can generate more solid waste for disposal. 7. Sub-sample the chromous chloride-sediment slurries for radioassay of unreduced ~S-sulfate (typically a 10-100pl volume is sufficient); decant the remaining slurry volume into waste jugs; rinse the flasks with tap water three times and decant the rinses into waste containers; the boiling flasks should then be suitable for reuse. Assaying the chromous chloride solution for radioactivity facilitates an accurate estimate of the amount of radioactivity added to a given sample. However, this may not prove necessary if the amount of added radioactivity is otherwise accurately known (e.g. by assaying aliquots of stocks injected into sediments; see 'introduction of ~SO~-~ into samples, incubation and termination' earlier in this section). Note: prior to replacing the scintillation vial traps for a new distillation, the Teflon tubing immersed in the traps should be wiped with a tissue to prevent sample carryover. Tissues should be placed in solid radioactive waste containers.
8. Reduction and trapping efficiencies can be estimated by distilling a finely ground commercially obtained pyrite standard (e.g. Alpha Aesar Company), and assaying the amount of sulfide contained in the zinc acetate traps. Combined reduction and trapping efficiencies typically exceed 90%. 9. Rates of sulfate reduction are calculated using the preceding formula, the sum of ~'S distilled (corrected as necessary for trapping and 495
¢-
~E o~
distillation efficiencies), the a m o u n t of ~SO~~ added (see step 7) and the concentration of sulfate in the samples. The latter term can be obtained readily using a turbidimetric assay for porewaters obtained by dialysis, centrifugation or squeezing from marine and some freshwater sediments (see Tabatabai, 1974; King and Klug, 1982; King, 1988). When sulfate concentrations are relatively low (<50 BM), HPLC methods with ion conductivity detection are preferable (Jorgensen and Bak, 1991). Sulfate concentrations measured per unit volume of porewater (i.e. in molar units) must be converted to a concentration per volume of sediment (e.g. ~tmol cm ~) using empirically determined water content data. Rates are typically expressed as ~tmol SO~2 reduced cm 3 d ', and depth profiles are typically integrated to yield units of mmol m : d ~.
Solutions 1. Mercuric chloride: 0.1 M in 1 M HC1. 2. Chromic chloride: 1 M in 1 M HC1. Chromous chloride is prepared as follows: 1. Immerse 100-200 g Zn metal (granular, 20-30 mesh size) in a solution of mercuric chloride and mix gently. After the metal acquires a bright metallic sheen, decant and save the mercuric chloride. Wash the Zn with tap water, transferring the first three rinses to containers for mercuric waste. Minimize exposure of Zn to the atmosphere during these washes. 2. Prepare 2-3 1 of chromic chloride in a large (4-6 1) Erlenmeyer flask; transfer the Zn to a second flask and add the chromic chloride. Stir until the solution turns deep blue from deep green. Do not seal the flask as it evolves hydrogen. Decant the chromous chloride and store in a sealed bottle. The solution remains stable for several weeks, or until it appears blue-green. 4. Wash and save the Zn metal; store submerged under deionized water. .
Cautionary notes Extensive experience indicates that >90% of the sulfide mass and radiolabel distilled from most sediments precipitates in the first of the three zinc acetate scintillation vial traps, with the remainder in the second trap. As a result, the third trap may be left in place as a safety measure, and changed intermittently. However, the distribution of mass and radiolabel should be checked empirically by each investigator. Formation of a white zinc sulfide precipitate provides a quick, convenient qualitative index of distribution. Investigators should plan in advance for storage and disposal of potentially large volumes of solid and liquid radioactive waste generated during sulfated reduction assays. Though the levels of radioactivity are typically low, they often exceed guidelines for sink disposal and in any
496
case, acidic chromous chloride requires specific disposal as a potential chemical hazard. Where space permits, liquid and solid wastes can be stored until radioactivity has decayed sufficiently for the material to be treated as chemical waste. Finally, investigators should exercise considerable caution when handling toxic metal solutions (chromous and mercuric chlorides). Amalgamation of zinc for chromium reduction produces a volatile form of mercury (elemental mercury) to which exposure should be eliminated. Likewise, amalgamated zinc should be stored so as to avoid any mercury volatilization into the lab environment. Even with quantitative radiosulfide precipitation in the zinc acetate traps, sediment distillations should be conducted with the distillation apparatus in a fume hood.
M O D I F I C A T I O N S OF T H E S I N G L E - S T E P PROCEDURE The single-step procedure may be modified to facilitate fractionation of reduced sulfur into various operationally defined pools (King, 1988). Distillation with acid only (1 N HC1 or H~SO~) yields a so-called acidvolatile sulfide fraction corresponding to poorly crystalline iron monosulfides (FeS). Subsequent to such a distillation, sediments can be collected by filtration and extracted with carbon disulfide to determine levels of elemental sulfur. After evaporating the CS~ to dryness, the sediment residue can be subjected to a chromous chloride distillation to yield estimates of pyritic sulfur. However, in view of the isotopic exchange that occurs among various sulfur pools (Fossing et al., 1992), there is usually little to be gained from multi-step processing for sulfate reduction rates. On the other hand, valuable assessments of -'~S'reduction and transformations of other sulfur species (e.g. thiosulfate) can be facilitated by appropriate modifications of the distillation described above (Troelsen and J~rgensen, 1982; Jorgensen and Bak, 1991; Elsgaard and Jorgensen, 1992). For example, an acid distillation followed by a chromous chloride distillation can be used to estimate radiosulfides formed from ~S' and label in '~S", "~S-thiosulfate, ~>S-polysulfide and ~S-sulfite phases, respectively. Of course, due consideration must be given to the possibility of elemental sulfur formation resulting from ferric iron released during the acid distillation step.
References Berner, R. A. (1964). An idealized model of dissolved sulfate distribution in recent sediments. Geochim. Cosmochim. Acta 28, 1497-1503. Canfield, D. E. (1989). Sulfate reduction and oxic respiration in marine sediments: implications for organic carbon preservation in euxinic environments. Deep-Sea Res. 36, 121-138.
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I,,, , D
"o
Capone, D. G. and Kiene, R. G. (1988). Comparison of microbial dynamics in marine and freshwater sediments: contrasts in anaerobic carbon metabolism. Limnol. Oceanoy,r. 33, 725-749. Cline, J. D. (1969). Spectrophotometric determination of hydrogen sulfide in natural waters. Limnol. Oceallogr. 14, 454-458. Elsgaard, L and Jorgensen, B. B. (1992). Anoxic transformations of radiolabeled hydrogen sulfide in marine and freshwater sediments. Geochim. Cosmochim. Acta 56, 2425-2435. Fenchel, T., King, G. M. and Blackburn, T. H. (1998). Bacterial Biogeochemistry: An Ecophysiological Analysis of Mim, ral Cycling. Academic Press, New York. Fossing, H. and Jorgensen, B. B. (1989). Measurement of bacterial sulfate reduction in sediments: evaluation of a single-step chromium reduction. Biogeochem. 8, 205-222. Fossing, H., Thode-Andersen, S. and Jorgensen, B. B. (1992). Sulfur isotope exchange between S-35 labeled inorganic sulfur compounds in anoxic sediments. Mar. Chem. 38, 117-132. Frtind, C. and Cohen, Y. (1992). Diurnal cycles of sulfate reduction under oxic conditions in cyanobacterial mats. Appl. Environ. Microbiol. 58, 70-77. Giray, C. and King, G. M. (1997). Effect of naturally-occurring bromophenols on sulfate reduction and ammonia oxidation in sediments from a Maine mudflat. Aquatic Microbial Ecol. 13, 295-301. Hansen, K., King, G. M. and Kristensen, E. (1996). Impact of the soft-shell clam, Mya arenaria burrows on sulfate reduction in an intertidal sediment. Aquatic Microbial Ecol. 10, 181-194. Howarth, R. W. and Giblin, A. E. (1983). Sulfate reduction in the salt marshes at Sapelo Island, Georgia. Limnol. Oceanogr. 28, 70-82. Howarth, R. W. and Merkel, S. (1984). Pyrite formation and the measurement of sulfate reduction in salt marsh sediments. Limnol. Oceanoy,r. 29, 598-608. Howarth, R. W. and Teal, J. M. (1979). Sulfate reduction in a New England salt marsh. Limnol. Oceanogr. 24, 999-1013. Howes, B. L., Dacey, J. W. H. and King, G. M. (1984). Carbon flow through oxygen and sulfate reduction pathways in salt marsh sediments. Limnol. Oceanogr. 29, 1037-1051. Ivanov, M. V. (1956). isotopes in the determination of the sulfate-reduction rate in Lake Belovod. Microbiologiya 25, 305 309. Jorgensen, B. B. (1978a). A comparison of methods for the quantification of bacterial sulfate reduction in coastal marine sediments. I. Measurements with radiotracer techniques. Geomicrobiol. J. 1, 11-27. Jorgensen, B. B. (1978b). A comparison of methods for the quantification of bacterial sulfate reduction in coastal marine sediments. II. Calculation from mathematical models. Geomicrobiol. J. 1, 29-47. Jorgensen, B. B. (1982). Mineralization of organic matter in the sea bed - the role of sulphate reduction. Nature 296, 643-645. Jorgensen, B. B. and Bak, E (1991). Pathways and microbiology of thiosulfate transformations and sulfate reduction in a marine sediment (Kattegatt, Denmark). Appl. Environ. Microbiol. 57, 847-856. King, G. M. (1983). Sulfate reduction in Georgia salt marsh soils: an evaluation of pyrite formation with ~Te and ~S tracers. Limm~l. Oceam~gr. 28, 987-995. King, G. M. (1988). The dynamics of sulfur and sulfate reduction in a South Carolina salt marsh. Limnol. Oceam)gr. 33, 376-390. King, G. M. (1990). Effects of a d d e d manganic and iron oxides on sulfate reduction and sulfide oxidation in intertidal sediments. FEMS Microbiol. Ecol. 73, 131-138.
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King, G. M. and Klug, M. J. (1982). Comparative aspects of sulfur mineralization in sediments of a eutrophic lake basin. Appl. EHviron. Microbiol. 44, 1406-1412. King, G. M., Howes, B. L. and Dacey, J. W. H. (1985). Short-term endproducts of sulfate reduction: formation of acid volatile sulfides, elemental sulfur, and pyrite. Geochim. Cosmochim. Acta. 49, 1561-1566. King, G. M., Carlton, R. G. and Sawyer, T. E. (1990). Anaerobic metabolism and oxygen distribution in carbonate sediments of a submarine canyon. Mar. Ecol. Prog. Ser. 58, 275-285. Martens, C. S. and Berner, R. A. (1977). Interstitial water chemistry of Long Island Sound sediments. I. Dissolved gases. Limnol. Oceatzoyr. 22, 10-25. Skyring, G. W. (1987). Sulfate reduction in coastal ecosystems. Geomicrobiol. J. 5, 374. Tabatabai, M. A. (1974). Determination of sulfate in water samples. Sulphur IHst. J.
10, 11-13. Troelsen, H. and Jorgensen, B. B. (1982). Seasonal dynamics of elemental sulfur in two coastal sediments. Estuar. Coast. Shelf Sci. 15, 255-266. Westrich, J. T. (1983). The consequences and controls of bacterial sulfate reduction in marine sediments. PhD dissertation, Yale University, New Haven, Connecticut. Zhabina, N. N. and Volko~; I. I. (1978). A method for determining various sulfur compounds in sea sediments and rocks. In: Ei~viromnelltal Biogeochemistry and Geomic~vbiolG~y (W. E. Krumbein Ed.), Voi. 3, pp. 735-745. Ann Arbor Science, Ann Arbor, Michigan.
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List of suppliers
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500
25 Nitrogen Fixation and Denitriftcation Douglas G Capone' and Joseph P Montoya ~ 'Wrigley Institute for EnvironmentalStudies & Dept. of Biological Sciences,University of Southern California, LosAngeles,CA 90089-03 7 I, USA;~Schoolof Biology,Georgia Institute of Technology, Atlanta, GA 30332-0230, USA
CONTENTS Introduction N z fixation DenJtritqcation "o
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INTRODUCTION The marine N cycle is an area of ever increasing research focus. The importance of the marine N cycle as a modulator for oceanic primary production and of the C cycle in marine ecosystems is well recognized (Capone, 2000). Compared to many other biogeochemical cycles, the N cycle is relatively complex, with biologically important species of N occurring in gaseous, dissolved and particulate phases, in inorganic and organic forms, and at a range of oxidation states. The natural microbiota are responsible for most of the key biological transformations of the N cycle and thereby contribute to its complexity. New pathways and nuances of the cycle continue to be discovered. Approaches to study the microbiological aspects of the marine N cycle are many and varied and new methods are constantly evolving. At the broadest level, our approaches to studying the marine N cycle can be regarded as either observational or experimental. Many of our insights and much of our current understanding into the role of biological transformations in the N cycle are grounded in geochemical observations over various spatial and temporal scales of the distributions of key nitrogenous species (Capone, 2000). However, inferences of pathways and fluxes derived from species concentrations and patterns typically only provide an estimate of the net result of multiple interacting processes. More recently, the application of the natural abundance of stable isotopes of N has greatly enhanced the utility of such observational approaches (Peterson and Howarth, 1987). Small differences in the ratio of
METHODS IN MICROBIOLOGY, VOLUME 30 ISBN ()-12 521530-4
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the stable isotopes of nitrogen, ~N and '~N, in different pools of N arise through isotopic fractionation associated with biological processes. As a result, the major biologically active pools of N in natural systems frequently have distinct isotopic signatures which can be exploited as an in situ tracer for studying the pathways of flow of nitrogen through whole ecosystems. In parallel with observational studies, experimental studies using natural and cultured microbial populations have greatly expanded our understanding of microbially mediated N transformations. Such experimental approaches of the N cycle have allowed qualitative confirmation of inferences and hypotheses developed using observational and classical microbiological methods. For example, nutrient measurements coupled to experimental biology (e.g. by sample isolation and incubation) first allowed us to directly quantify and compare rates of N transformations within and among various environments (Capone, 1996). Note, however, that the time-course of change in a particular chemical pool over time must be interpreted with some care since complementary or opposing processes may produce and consume a particular compound simultaneously (e.g. nitrification produces NO~ while denitrification consumes NO3 ), making it difficult to isolate the activity of an individual process. If these interacting processes occur at comparable rates, then time courses from incubations may reveal little net change in the pool of interest (i.e. steady state) despite significant fluxes in and out of that pool. The enzymes mediating various N transformations (e.g. assimilatory NO~ reductase, glutamine synthetase) have been characterized and assays developed to estimate their activity in natural populations of marine microbes (Falkowski, 1983). Inhibitors of particular pathways may be used to block one process, thereby revealing gross rates of other processes (Oremland and Capone, 1988). Direct tracer methods provide less ambiguous means to quantify mass fluxes directly through specific pathways of biological N transformation (Harrison, 1983). While N does have a radioisotope, '~N, its very brief halflife and the sophisticated facilities necessary to generate it, preclude its use in most field applications (Capone, 1996). In contrast, the stable isotope of N, '~N, is available in a variety of forms useful for determining rates of specific pathways in the N cycle. Recently, immunological and molecular methods have provided sensitive means to detect proteins and genes involved in N cycle transformations (Ward, 1995; Zehr and Capone, 1995). With improvement in reverse transcriptase (RT) methods, analysis of genes, transcripts (mRNA) and gene products (proteins), it may soon be possible to comprehensively analyze the induction and regulation of the various enzymes responsible for N transformations. For the purposes of this presentation, we will focus on the most widely used enzyme-based and direct tracer procedures for two key microbial processes in the N cycle, N, fixation and denitrification. A variety of other procedures appropriate to various situations are available and are summarized in Capone (1996, 2000) and Seitzinger et al. (1993). 502
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N2 F I X A T I O N
Principle Biological nitrogen fixation is the enzymological capacity of certain bacteria and Archaea to convert gaseous dinitrogen to ammonium on a pathway to amino acid synthesis (Postgate, 1998). In marine systems, the greatest attention has focused on diazotrophic cyanobacteria, largely because of their quantitative importance in supplying new N to the upper ocean (Capone et al., 1997; Capone and Carpenter, 1999). However, N, fixation by many other physiological groups is recognized (Capone, 1988; Zehr et al., 2000) and may also be quantitatively significant in the oceanic N budget. Nitrogenase activity is routinely determined in many laboratories using the C~H_~reduction procedure (Stewart et al., 1967; Hardy et al., 1968; Capone, 1988). C~H~is a substrate analog of N~ (its natural substrate) (Postgate, 1998). When added in relatively high concentrations (e.g. pC~H~ > 0.1 atm), C~H~ is preferentially reduced by nitrogenase to C2H~ which is easily quantified by flame ionization gas chromatography. Small gas chromatographs are inexpensive, durable, highly portable, and may be set up at remote sites with limited logistical support. Alternatively, experimental systems can be exposed to ~N~ enriched to levels far above its natural abundance (Montoya et al., 1996). The flux of labeled nitrogen into the product (e.g. cell N or NH/) over appropriate periods is determined using emission or isotope ratio mass spectrometry. The two approaches are conceptually similar and will be presented together.
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Sealable assay vessels with very low gas permeability and means for headspace gas sampling and withdrawal.We routinely use glass serum vials available in a range of sizes from about 7 to 150ml with silicone rubber crimp-seal closures. For more fastidious anaerobes, butyl stoppers are available. Larger (e.g. 250 ml) bottles, with Teflon faced, butyl screw-top closures are available from Wheaton. For larger or unusual samples, custom vessels of glass, plexiglass or flexible material (e.g. Saran wrap) with fitted injection port may be fabricated and used. • Gas-tight syringes (e.g. Hamilton) of appropriate volume for (I) injection of acetylene or 'SN2 (usually 0.1 to 10 ml): for the 'SN~ we generally employ syringes equipped with an opening-closing valve as well), and (2) sub-sampling of the headspace of experimental vessels for chromatographic analysis in the C2H2 reduction procedure.We routinely use 00 pl syringes for the latter.
For the CzH ~ reduction procedure • A source of clean acetylene. We use either instrument grade C2H 2 with a sparge through water (to remove traces of acetone: see below), or acetylene generated on site by reaction of water and CaC 2. For the latter, we employ a small (e.g. 150 ml) side arm flask with a piece of tygon tubing attached to the
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side arm and terminated into a I cc syringe barrel (plunger removed and distal end cut off. A long (>_ 1.5", 20 g) needle is attached to the syringe barrel and the needle inserted into a sealed glass serum vial, typically about 20 ml, twothirds filled with deionized water.The needle should penetrate the water.An excurrent 18g needle penetrating to the headspace is used to provide a sampling port for the gas tight needle used for C~H2 addition. Pellets of CaC2 are added to the flask, the flask sealed with a black rubber stopper and the C2H2 formed allowed to continuously bubble through the serum vial. Gas samples are taken by inserting a gas-tight syringe with a 22 g needle through the center of the 18 g excurrent into the headspace. CaC~ is added as necessary. C2H ~ may be generated in the laboratory and stored in a vessel (e.g. a football bladder works quite well, Paerl, personal communication) for subsequent use in the field. Flame ionization gas chromatograph (FID/GC) fitted for detection of light hydrocarbons and particularly for baseline separation of ethylene and acetylene in that order. We routinely use 2 m x I/8" columns of HaySep A (80/100 mesh) available from most chromatographic suppliers. Peak heights are quantiled either using a strip chart recorder or, preferably, a simple integrator.We have found better precision for the sharp peaks formed operating in the in the peak height (rather than area) mode.Very low levels of C2H~ production may be achieved using laser photoacoustics (Zuckermann et al., 1997). Standards for C~H~.We generally prepare standards from the pure gas by serial dilution in sealed 250 ml glass bottles or use certified, commercially available low pressure l00 ppm (v/v) standards obtained from AIItech or Scott. For the latter, we modify the plastic pressure valve, removing the stainless steel needle from the plastic tube such that we can insert and fill our 100pl syringe directly from the can. It is useful to periodically ensure that the FID response is linear over the range of concentrations of C~H~ experimentally encountered. For the 'SN 2 uptake procedure
• A source of 'SN2 obtained commercially (e.g. Cambridge Laboratories or Isotec) or generated from oxidation of 'SNH~+ salts (Burris, 1974). Commercial 99 atom % 'SN2 is available in small (e.g. 0.5 I) steel cylinders with total volumes of about 4 I.We have fabricated a small sampling manifold for attachment to the cylinder; this manifold is equipped with a low pressure gauge, syringe port and low dead space that allows pressurization of the valve assembly, and withdrawal of discrete volumes into gas-tight needles with shutoff valve, as needed for individual assays. • Filtration apparatus with which to collect biological samples at incubation termination. We generally collect particulate material on pre-combusted (450°C)Whatman (or equivalent) 25 mm GF/F (nominal pore size 0.7 pro) or GF/C (nominal pore size 1.4 pm) glass fiber filters. • An isotope ratio mass spectrometer (IRMS) optimized for sampling masses 28 and 29 and equipped for either on-line combustion of particulate samples or for introduction of gas samples from samples converted off-line to N 2. Alternatively, one may use an emission spectrometer (Fiedler and Proksch, 1975).
504
Assay I: CzHz reduction procedure •
•
•
•
•
For marine samples, we typically employ a water phase to gas phase ratio of about I:1. Place biological sample to be analyzed in assay vessel. Hence, for unconcentrated or concentrated samples of seawater, fill approximately half the container. For macroparticulate samples (e.g. macroalgae, cyanobacterial mat), we generally bathe the sample in 0.2-pm-filtered ambient seawater (it is useful to determine the inorganic N content of the water phase). Marine sediments may be assayed as intact cores, or as sediment-seawater slurries, depending on the intent of the experiment. Sample vials are sealed with a septum and crimp-seal closure. Depending upon the type of sample and the anticipated type of diazotrophic flora, attention must be paid to 02 exposure as many diazotrophs are highly sensitive to 02 . Hence, an experimental setup minimizing or avoiding exposure to O 2, for example in a glove bag under N 2 or in an anaerobic hood, may be indicated. Once the vial is sealed and any necessary modification of the assay environment is made (see below), the C2H2 reduction assay is initiated by injection of sufficient C~I-I2co achieve a pC2H2 of at least 0. I atm. Samples are incubated under environmental conditions appropriate for the samples. Experiments may be run over a discrete time period and activity terminated by use of a chemical or physical poison (e.g. by addition of T C A or by rapid freezing) until headspace analysis. However, we generally sample over a time course, periodically removing 100 IJI samples for immediate analysis of C2H4 and C2H 2 by FID/GC. (Alternatively, gas phase samples may be removed and injected into Vacucainers", or evacuated serum vials, for analysis at a later date.) Depending on the level of nitrogenase activity, C2H 4production may be manifested in minutes (e.g. cyanobacterial mats) or hours to days (organicrich surficial sediments). C2H4 is quantifed by comparison with standards (see above). C2H 4concentrations are scaled to the volume of the gas phase of the assay vessel and corrected for C2H4dissolved in the water phase with consideration of salt and temperature effects on the solubility coefficient (Capone, 1993).The calculations used are:
C2H4formed -
Peak htsample Peak
xConc, std(nmol ml I)xGPVxSC
(25.1)
htstanclar d
where GPV = gas phase volume of assay vessel in ml, SC = solubilty correction for C2H4in the aqueous phase. SC is calculated according to Flett et al. (I 976)
SC-- M/X = I + (o0 (A/B) where M X
A B
(2s.2)
= total volume of C2H4 produced, = volume in gas phase, = Bunsen coefficient for CzH4 at the appropriate temperature and salinity, = volume of the aqueous phase, = volume of the gas phase (or GPV).
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We generally derive our rate estimate by linear regression analysis of the total C2H4 produced as a function of time, ideally using data obtained closest co initiation of the experiment as the best approximation of the initial rate. Depending on the sample, rates expressed as moles C2H4 per unit time are normalized to volume (e.g. per ml of seawater or sediment assayed), biomass (as particulate N, dry weight, or for algal samples, chlorophyll) or crosssectional area (e.g. assaying discrete sediment cores or algal mats of known cross-section or by scaling biomass assays by volumetic density and depth integrating).The measured rates of C2H ~ reduction are then converted to N~ fixation rates using either a standard reduction ratio (see Applications/ Quantification) or a reduction ratio determined by parallel incubations with C~H~and ~N:.
Assay I1: '~Nz uptake • To simplify the calculation of substrate isotopic enrichment, samples are placed in assay vessels that are filled completely with either the sample itself (e.g. seawater, or seawater concentrates) or for particulate samples, with 0.2IJm-filtered ambient seawater and sealed. Separate, comparable samples should be taken for initial particulate N concentrations, treated as below. • As mentioned above, depending on the system, care may be necessary to minimize or fully avoid exposure to 02. • Exogenous substrate or inhibitors can be added at this point, or samples preincubated as appropriate (see below). • The assay is initiated by the addition of a small bubble of ~SN2 gas through the septum (e.g. 0.5 ml in a 250 ml vessel, which yields enrichment of about 12-13 atom %, depending on temperature and salinity), usually with a gas-tight syringe equipped with an opening-closing valve.We routinely fill the syringe at a pressure slightly above ambient, then vent the syringe to bring it to I arm pressure before adding the tracer gas to the incubation vessel; this procedure ensures that the syringe is completely filled with ~SN2 at I arm pressure. An equivalent volume of solution is removed to equalize the pressure across the septum. • Samples are incubated under conditions of light and temperature as appropriate for the system being examined for several (e.g. 3 to 24 h, depending on anticipated activity) hours. • The experiment is terminated by unsealing the container and either filtering the contents with gentle vacuum (<25 cm Hg) onto pre-combusted glass fiber filters (planktonic material), or by removing and rinsing the macroparticulate matter. Samples are either immediately dried at 60°C and stored desiccated, or frozen to less than -20°C until dried. • Samples are analyzed for isotopic ratio of N by IRMS (Montoya et al., 1996) or emission spectrometry (Fiedler and Proksch, 1975). • N2 fixation is calculated from the increase in 'SN content of the sample relative to isotopic enrichment of the substrate pool:
N2fixation (moles N 2 t I) =
APpN final -- APpN-initial PNr~naI (mol) I × - At APN2 - APpN_initiaI × 2
506
(25.3)
where: AP = atom % 'SN of the sample (e.g. PN~,,~,)or substrate (i.e. N2) pool. Although AP........, can often be approximated using the global average abundance of 'SN (0.366 atom %), a direct measurement is necessary to quantify low rates with high precision. The atom % and mass of the PN pool are determined by mass or emission spectrometry and elemental analysis, respectively while the '~N enrichment of the substrate (N~) is calculated from the dissolved mass of N~ in the assay system based on the solubility of N~ at the assay temperature and salinity (Weiss, 1970), and the mass and isotope ratio of the N, addition (Montoya et al., 1996). As for C~H: reduction, rates are generally normalized to volume (per ml or 1), biomass (per unit N, mass, or chlorophyll) or cross-sectional area (m-3.
Troubleshooting Tile C2H~ reduction method is a relatively sensitive, simple and straightforward procedure. However, there are potential confounding factors. One should take care to ensure that there are no contaminants when using commercial C:H: (Hyman and Arp, 1987). C~He is itself an inhibitor of actual N~ fixation and, hence, extended assay times may cause responses to N starvation in the system under examination including lags a n d / o r accelerating rates. Lag periods may be observed which can be a result of physical (C~H~ solution) or biological (enzyme induction or diazotroph proliferation) factors. The direct N, fixation assay is very sensitive and especially well suited to characterizing the rate of fixation in dilute suspensions of diazotrophs. In field situations where multiple rates are being measured in parallel experiments, the major potential source of difficulty with this method is the possibility of contamination of the sample with '~N-enriched materials associated with other tracer measurements. Dedicated filtration and sample handling equipment should be used for this assay, and the experimental apparatus should be physically separated from the site of highlevel tracer experiments. Similarly, these samples should be analyzed with others of similarly low enrichment in order to minimize the potential effect of sample carryover during the mass spectrometric analysis.
Application Controls A variety of controls are recommended when first examining a new system for either assay. Abiological controls (e.g. sterilized, chemically inhibited or without addition of biological material) are appropriate for both. C~H~-free controls for the C:H: reduction procedure provide a check on natural C~_H~production. C:Ha-enriched samples, without C~H~ allow detection of hydrocarbon oxidizers that might be present. C~H~may itself be used as a control for '~N~uptake experiments.
507
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Gas:water phase ratios As pointed out by Flett et al. (1976), the lower the gas phase to aqueous phase ratio, the potentially more sensitive the C~H~ reduction assay system. However, in many cases, maximal sensitivity is not required, and a larger gas phase ratio allows repetitive sampling of small gas volumes over a time course without introducing a significant negative pressure. Furthermore, a very small gas phase volume maximizes the concentration of the background C,H~ peak present in most sources of c2Iq:, thereby compromising the increase in sensitivity.
Experimental manipulation Once sealed and before initiation of the assay by addition of C,H~ or '~N~ the atmosphere in the samples may be modified (e.g. O~ tension), samples may be pre-incubated under various conditions of light, temperature, or inorganic or organic substrates or metabolic inhibitors added to the experimental system.
Quantification If one's intent is to estimate quantitatively N~ fixation in situ, this will be derived directly from the ~N 2 assay. For C,Ho-reduction-based experiments, N~ fixation rates are calculated using a reduction ratio (C2H~reduction:N~ reduction) that is either measured directly in parallel incubations or assume to conform to a standard biochemical stoichiometry. The theoretical reduction ratio is 3:1, which is based on the number of reducing equivalents required by nitrogenase to reduce N~ to ammonium (6) versus C2H2 to C2H 4 (2). However, a ratio of 4:1 may be more appropriate if one allows for the fact that under N:-fixing conditions, approximately one mole of H~ is evolved per mole of N: fixed (total of eight reducing equivalents). This 'hydrogenase' activity of nitrogenase is greatly reduced under C2H2-reducing conditions, resulting in a relatively greater yield of C~H4 (Postgate, 1998). It is generally advised, however, to compare directly and thereby calibrate, these two methods for particular systems. One important distinction that should be pointed out between these two approaches, however, is that whereas C2H~ reduction can (given a conducive environment) potentially assess the activity of all extant nitrogenase activity in a system, I~N~uptake is generally only measured by its flux into the particulate fraction. Any recently fixed N which is released in soluble form (e.g. as ammonium, amino acids) will not be quantified unless specific procedures are implemented to sample those other target pools (Glibert and Bronk, 1994).
4~4~4~4~4~4~ D E N I T R I F I C A T I O N Principle Denitrification can be measured conveniently in many microbial systems by addition of C2H2 which inhibits N20 reductase, the last step of the
508
denitrification pathway (Yoshinari and Knowles, 1977; Sorensen, 1978). The presence of C~H~ results in the accumulation of N~O which can be selectively and very sensitively detected by electron capture gas chromatography (Capone, 1996). Many of the procedures and conceptual design of denitrification experiments parallel those of the C~H~ reduction procedure. Indeed, one may use the same assay to monitor for both activities (Capone and Taylor, 1980; Joye and Paerl, 1993). As for nitrogen fixation, denitrification can also be measured directly by the introduction of enriched 15-N nitrate (or nitrite) into a system, with the determination of progressive enrichment of 15N in the N~ pool. C o m m o n equipment and reagents • As above, gas-tight syringes and sealed assay vessels.
For the C2H 2 blockage procedure • A clean source of C2H 2. • Gas chromatograph equipped with electron capture detector and configured for separation of 02, COs, N20 and H2S.We typically employ a HaySep Q column, I/8" × 2 m, 80/100 mesh. Integrator or strip chart recorder for peak height analysis. • Standards for N20. One may make serial dilutions from pure standards, or obtain certified standards from chromatography suppliers. Depending on the concentration range of N20 anticipated, standards over the range 0. I to 100 ppm (v/v) are suggested.
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For the ~SN2 tracer procedure • A source of 'SN-NO3 obtained commercially (e.g. Cambridge Laboratories or Isocec).We typically prepare a concentrated stock solution (e.g. I raM) for addition to the incubation vessel.The tracer stock solution is sparged with He to remove 02, then kept in a sealed vessel under an oxygen-free headspace. • Large volume gas-tight syringes for sampling the incubation vessel headspace or liquid at the end of the experiment. • A vacuum line for extraction and purification of the N 2 evolved during the incubation. • An isotope ratio mass spectrometer optimized for sampling masses 28 and 29 and equipped for either continuous-flow analysis of gas samples injected directly into a carrier stream of He, or with a dual inlet for the introduction of gas samples from samples of N 2extracted and purified with a vacuum line. Alternatively, one may use an emission spectrometer (Fiedler and Proksch, 1975).
Assay I: CzH2 blockage procedure • As for C2H 2 reduction samples, we typically employ a water phase to gas phase ratio of about I:1. Place biological sample to be analyzed in assay vessel. Hence, for unconcentrated or concentrated samples of seawater, fill
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•
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•
•
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approximately half the container. (As mentioned, one may simply measure N20 in assays set up for C2H 2 reduction.) Sediments are often examined for denitrifying activity.They may be assayed as intact cores extruded into a larger assay vessel, or as sediment-seawater slurries, depending on the intent of the experiment. As for nitrogenase activity, attention needs to be paid to O2 concentration in assays as many denitrifiers are facultative anaerobes, synthesizing the enzymes for denitrification only in the absence of O2 (Knowles, 1982). Hence, an experimental system that either minimizes or avoids exposure to 02, for example in a glove bag under N2 or in an anaerobic hood, may be indicated.Alternatively, 02 may be removed by sparging with N2 after the system is sealed. Once the assay vessel is sealed and any desired modification of the experimental environment is made (see below), the assay is initiated by injection of sufficient C2H2to achieve a pC~H2 of at least 0. I arm. Samples are incubated under environmental conditions appropriate for the samples. For sediments, as an alternative, cores may be retained in mini-core barrels (e.g. three 50 cc syringes with the front (needle) end cut off, the plunger retained and the open end sealed with a rubber stopper with a recess. Solutions of deoxygenated seawater saturated with C~H 2 are then injected through the stopper into the core. Experiments may be run over a discrete time period and activity terminated by use of a chemical or physical poison (e.g. by addition of T C A or by rapid freezing) until head space analysis. However, we generally sample over a time course, periodically removing 100 lul samples for immediate analysis of N2© by ECD/GC (Alternatively, gas phase samples may be removed and injected into Vacutainers ~, or evacuated serum vials, for analysis at a later date.) For intact core assays,the cores are extruded into an appropriate vessel at the end of the incubation period, rapidly sealed, disaggregated by shaking and the N~O formed is allowed to equilibrate with the gas phase. Subsamples of the gas phase are collected and analyzed for N20 content as above. N20 is quantifed by comparison with standards (see above). N20 concentrations are scaled to the volume of the gas phase of the assay vessel and corrected for N20 dissolved in the water phase with consideration of salt and temperature effects on the solubility coefficient (Capone, 1993).The calculations used are directly analogous to those for C2H2 production (Equations (25.1) and (25.2)). Rate estimates are derived as for the C2H2 reduction procedure.
Assay Ih 'SN tracer procedure • Addition of 'SN-labelled inorganic substrate to an experimental system allows direct quantification of the rate of denitrification as a flux of tracer from the dissolved into the gaseous pools.This experimental approach can be generalized to allow quantification of multiple fluxes in parallel incubations with different labeled substrates (e.g. 'SNO3 for denitrification, 'SNH4+ for coupled nitrification-denitrification). The sample must be incubated in a gas-tight container which may contain a head space for gas sampling, or may be filled completely with the liquid phase in studies of pelagic denitrification.The presence of a headspace in the incubation vessel simplifies the final gas assay and
510
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allows for time course sampling from the head space, similar to the gas chromatographic procedures. However, the presence of a gas phase may not be practical for experiments with pelagic denitrifiers where introduction of a headspace may alter the gas composition of the experimental system or may add to the ambient pool of N2, thereby diluting the signal derived from production of ~N-labelled N2. In such cases, the incubation is carried out in a completely full assay vessel, and the dissolved gases are extracted from a water sample drawn at the end of the incubation (e.g. by a headspace equilibration in vacuo). Sediments may be assayed as intact cores extruded into a larger assay vessel, or as sediment-seawater slurries, depending on the intent of the experiment. Here again, the introduction of a headspace may simplify the gas sampling procedure, albeit at the cost of reducing the experimental sensitivity (by adding to the ambient burden of N2) and/or altering the gas composition of the experimental system. As for the C2H2 block assay, the experimental system should be designed to minimize or avoid exposure to 02, or O2 may be removed by spargJng with He after the system is sealed. Since this experimental approach involves a measurement of the quantity of '~N moving into the gaseous pools, it is advantageous to minimize the burden of N2 gas present in the system, which can also be accomplished by sparging with He orAr. Once the assay vessel is sealed and any desired modification of the experimental environment is made (see below), the assay is initiated by addition of sufficient 'sNO3 to achieve a source enrichment of about I 0 atom %. Samples are incubated under environmental conditions appropriate for the system. Experiments should be run over a discrete time period and either sampled directly or activity terminated by use of a chemical or physical poison (e.g. by addition of HgCI2) until a sample can be withdrawn for gas extraction and analysis. We typically use a large volume (10 ml) gas-tight syringe equipped with a valve to sample either the head space or the liquid phase of the incubation vessel. For intact core assays, the cores are extruded into an appropriate vessel at the end of the incubation period, rapidly sealed, disaggregated by shaking and the gases formed are allowed to equilibrate with the bull< solution, after which the gases are extracted as for a liquid sample. In some experimental systems, a gas sample can be collected for isotopic analysis directly from a headspace in the incubation vessel. For experiments conducted without a headspace (e.g. in studies of pelagic denitrification), the N2 can be isolated for isotopic measurement by transferring the water sample to a separate vessel for headspace equilibration (Emerson et al., 199 I). In either case, the isolated gases are dried, then N 2 can be isolated by cryogenic distillation using a vacuum line (Brandes and Devol, 1997), after which the isotopic composition of the N 2and/or N20 can be measured by dual-inlet isotope ratio mass spectrometry.Alternatively, the gases collected can be injected into a suitably modified elemental analyzer or gas chromatograph for on-line purification and continuous-flow isotope ratio mass spectrometry.The rate calculation is analogous to that for the Nrfixation assay (Equation (3)), with NO3 treated as the source pool (analogous to the ~SN2 pool in Equation (3)) and N2 treated as the target pool (analogous to the PN pool in Equation (3)).
511
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Troubleshooting Electron capture detectors, with their great sensitivity, can also encounter problems with interference from other compounds in the headspace eluting after N20 (e.g. C2H~, H2S, H~O) resulting in spurious peaks and drifting baselines. Many researchers employ chromatographic valving methods, such as a 'backflush to vent' to clean the column between analyses, or to use dual columns allowing one to flush while using the other. In many systems, a critical substrate for denitrification, NO~ may be in limited supply. Low NO~ concentrations can compromise the efficiency of the C~H, block (Koike and Hattori, 1978; Slater and Capone, 1989). The primary difficulty with the '~N-tracer assay for denitrification is to overcome the large burden of ambient No in order to measure the typically small fluxes of '~N into the gaseous pools. Where practical, this problem may be ameliorated by sparging the sample to reduce or remove the initial concentration of N,.
Application Many of the provisos for nitrogen fixation relating to controls and experimental manipulations apply equally to denitrification studies. Hence, for the C2H2 block procedure, killed controls are routinely carried out, and one should also determine the efficacy of the QH~block by measuring the disappearance of exogenously supplied N,O in the presence of CM~ In order to overcome problems of low NO,, many researchers often include experimental levels with a NO~ spike. Similarly, chloramphenicol is incorporated into assay media where one is interested in estimating in situ activity, in order to prevent the synthesis of new denitrifying enzymes (Joye and Paerl, 1993). However, chloramphenicol may interfere with extant enzyme (Wu and Knowles, 1995). Studies aimed at elucidating the factors controlling the rate of denitrification in a particular system may include addition of organic substrates (Brettar and Rheinheimer, 1992), manipulation of O: concentration, and alteration of the rate of NO~ supply. As mentioned above, it is often of interest to determine the relative importance of exogenous and in situ (i.e. nitrification) sources of NO~ supporting denitrification in surficial sediments. In addition to the direct tracer approach of adding '~NH, ~ and monitoring the flux of ~N to '~N~, a method referred to as isotope pairing has been used to estimate tile relative contributions of exogenous and internal sources of NO~ to denitrification (Nielson, 1992). Labeled, highly enriched '~NO, is added to nitrate-free water overlying a sediment core and the production of '~N~4N (mass 29) and '~N2 (mass 30) is measured over time. The relative proportions of denitrification resulting from '~NO~ (exogenous source) and '~NO, (internal sources, i.e. nitrification) can be directly calculated as a function of tile abundance of these dinitrogen isotopomers.
512
References Brandes, J. A. and Devol, A. H. (1997). isotopic fractionation of oxygen and nitrogen in coastal marine sediments. Geochim. Cosmochim. Acta 61, 1793-1801. Brettar, I. and Rheinheimer, G. (1992). Influence of carbon availability on denitrification in the central Baltic Sea. Limnol. Oceanoy,r. 37, 1146-1163. Burris, R. H. (1974). Methodology. In: The Biology qf N, Fixation (A. QuispeI, Ed.), p. 9. North-Holland, Amsterdam. Capone, D. G. (1988). Benthic nitrogen fixation. In: Nitrogelz Cycling in Coastal Marine Enviromnents (T. H. Blackburn and J. Sorensen, Eds), pp. 85-123. J. Wiley & Sons, New York. Capone, D. G. (1993). Determination of nitrogenase activity in aquatic samples using the acetylene reduction procedure. In: Hmutbook q[ Methods in Aquatic Microbial Ecolo~,,y (P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 621-631. Lewis Press, Boca Raton, FL. Capone, D. G. (1996). Microbial nitrogen cycling. In: Mamlal off E1n~iromnental Microbiology, Sectiotl IV: Aquatic Euviromnelzts (S. Newell and R. Christian, Eds), pp. 334-342. ASM Press. Capone, D. G. (2000). The marine nitrogen cycle. In: Marille Microbial Eeolo~,~y(D. K. a. R. Mitchell, Ed.). Wiley, New York. Capone, D. G. and Carpenter, E. J. (1999). Nitrogen fixation by marine cyanobacteria: Historical and global perspectives. Bull. Inst. OceaHogr. Monaco 19, 235-256. Capone, D. G. and Taylor, B. E (1980). Microbial nitrogen cycling in a seagrass community. In: Estuaril~e Perspectives (V. S. Kennedy; Ed.), pp. 153-161. Academic Press: New York. Capone, D. G., Zehr, J., Paerl, H., Bergman, B. and Carpenter, E. J. (1997). Trichodesmium: a globally significant marine cyanobacterium. Scie~lce 276, 1221-1229. Emerson, S., Quay, P., Stump, C., Wilbur, D. and Knox, M. (1991). O , At, N~, and :~Rn in surface waters of the subarctic ocean: net biological O~ production. Glob. Bio~?eochem. Cycles 5(1), 49-69. Falkowski, P. (1983). Enzymology of nitrogen assimilation. In: Nitro~,~eJl iH the Marille Em~iromneJlt (E. J. Carpenter and D. G. Capone, Eds), pp. 839-868. Academic. Fiedler, R. and Proksch, G. (1975). The determination of nitrogen-15 by emission and mass spectrometry in biochemical analysis. A review. Aiml. Chem. Aeta 78, 1 62. Flett, R. J., Hamilton, R. D. and Campbell, N. E. R. (1976). Aquatic acetylene-reduction techniques: solution to several problems. Cruz. J. Microbio[. 22, 43. Glibert, P. M. and Bronk, D. A. (1994). Release of dissolved organic nitrogen by marine diazotrophic cyanobacteria, Trichodesntium spp. Appl. E~nqron. Microbiol. 60, 3996-4000. Hardy, R. W. E, Holsten, R. D., Jackson, E. K. and Burns, R. C. (1968). The acetvlene-ethylene assav for N, fixation: laboratory and field evaluation. Plm~t Physiol. 43, 1185. Harrison, W. G. (1983). Use of isotopes. In: Nitrogel~ in the Marille E~nqromneut (E. J. Carpenter and D. G. Capone, Eds), pp. 763 807. Academic Press, New York. Hyman, M. R. and Arp, D. J. (1987). Quantification and removal of some contaminating gases from acetylene used to studv gas-ufilizing enzymes and microorganisms. Appl. DnqroJ1. Microb. 53, 298. Joye, S. B. and Paerl, H. W. (1993). Contemporaneous nitrogen fixation and denitrification in intertidal microbial mats: Rapid response to runoff events. Mar. Ecol. Pro~. Set. 94, 267 274.
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Knowles, R. (1982). Denitrification. Microbiol. Rev. 46, 43-70. Koike, I. and Hattori, A. (1978). Denitrification and ammonia formation in anaerobic coastal sediments. Appl. Enviroll. MicrobioL 35, 278-282. Montoya, J. P., Voss, M., Kaehler, P. and Capone, D. G. (1996). A simple, high precision tracer assay for dinitrogen fixation. Appl. Environ. Microbiol. 62, 986-993. Nielsen, L. P. (1992). Denitrification in sediments determined from nitrogen isotope pairing. FEMS Microbiol. Ecol. 86, 357 362. Oremland, R. S. and Capone, D. G. (1988). Use of specific inhibitors in microbial ecological and biogeochemical studies. Adv. Microbial Ecol. 10, 285-383. Peterson, B. J. and Howarth, R. W. (1987). Sulfur, carbon and nitrogen isotopes used to trace organic matter flow in the salt-marsh estuaries of Sapelo Island, Georgia. Limnol. Oceanogr. 32, 1195-1213. Postgate, J. R. (1998). The Flmdame~ttals o~ NitrogeJi Fixation. Cambridge University Press, Cambridge. Seitzinger, S. P., Nielsen, L. P. and Caffrey, J. (1993). Denitrification measurements in aquatic sediments: a comparison of three methods. Bi~geochem. 23(3), 147. Slater, J. M. and Capone, D. G. (1989). Nitrate requirements for acetylene inhibition of nitrous oxide reduction in marine sediments. Microb. Ecol. 17, 143-157. Sorensen, J. (1978). Denitrification rates in a marine sediment as measured by the acetvlene inhibition technique. Appl. EllviroH. Microbio/. 35, 301. Stewart, W. D. P., Fitzgerald, G. P. and Burris, R. H. (1967). lu situ studies on N~ fixation, using the acetylene reduction technique. Proc. Natl. Acad. Sci. USA 58, 2071. Ward, B. B. (1995). Functional and taxonomic probes for bacteria in the nitrogen cycle. In: Molecldar Ecology of Aquatic Microbes, I, pp. 73 86. Joint, NATO. Weiss, R. E (1970). The solubility of nitrogen, oxygen, and argon in water and seawater. Deep-Sea Res. 29, 459-469. Wu, Q. and Knowles, R. (1995). Effect of chloramphenicol on denitrification in Felxibacter canadellis and Pseudomonas deJfftrificmts. AFpl. El~vin~n. Microbiol. 61, 434-437. Yoshinari, T. and Knowles, R. (1977). Acetylene inhibition of nitrous oxide reduction and measurement of denitrification and nitrogen fixation in soil. Soil Biol. Biochem. 9, 177-183. Zehr, J. P. and Capone, D. G. (1995). Problems and promises of assaying the genetic potenial for nitrogen fixation in the marine environment. Microbial Ecol. 32, 1-38. Zehr, J. P., Carpenter, E. J. and Villareal, T. A. (2000). New perspectives on nitrogen-fixing microorganisms in subtropical and tropical open oceans. Tremls Microbiol. 8, 68-73. Zuckermann, H., Staal, M., Stal, L., Reuss, J., Hekkert, S., Harren F. and Parker, D. (1997). On-line monitoring of nitrogenase activity in cyanobacteria by sensitive laser photoacoustic detection of ethylene. Appl. Enviro~l. Microbiol. 63, 4243 4251.
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26 Optical Remote Sensing Techniques in Biological Oceanography W Paul Bissett', Oscar Schofield 2, Curtis D Mobley 3, Michael F C r o w l e y 2 and M a r k A Moline 4 'Florida Environmental Research Institute, Tampa, Florida, USA;~Coastal Ocean Observation Laboratory, Institute of Marine and Coastal Sciences,Rutgers University, New Brunswick, New Jersey, USA; 3Sequoia Scientific, Inc., Redmond,Washington, USA;4Biological Sciences Department, California Polytechnic State University, San Luis Obispo, California, USA
CONTENTS Introduction
Principle Hardware, software, and data Application Future directions
e, e(1) un
Conclusions .E remote sensor n. any instrument, such as a radar device or camera, that scans the earth or another planet from space in order to collect data about some aspect of it. - - r e m o t e - s e n s i n g adj., n. (Collins English Dictionary)
INTRODUCTION Light from the sun is the driving energy source behind all of the surface water biological processes. The radiant energy is harvested and stored as chemical energy through the process of photosynthesis providing the organic fuel for most of the oceanic food web. Single-cell marine phytoplankton are responsible for the majority of this energy conversion, and the growth of their organic biomass via autotrophic photosynthesis is referred to as p r i m a r y production (Parsons et al., 1984). Oceanic net ~ p r i m a r y production is about one-third of the global net primary production (Denman et al., 1996). The estimate of oceanic biomass and net p r i m a r y production has been revised u p w a r d s over the last two decades. M E T t l O D S IN MICROBIOLOGY, VOLUME 30 ISBN 0 - I 2 521530-4
C o p y r i g h t © 2001 A c a d e m i c Press Ltd All rights of r e p r o d u c t i o n in any form reserved
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This revision occurred in part because of the data stream provided by the first ocean color satellite sensor, the Coastal Zone Color Scanner (CZCS), and the scientific efforts of the NIMBUS-7 Experiment Team (NET) and many other ocean color scientists (Acker, 1994). As visible light enters the water column, the in situ constituents, including water itself, impact the light's directionality and color. In pure seawater, blue light (-430 nm) is least impacted by the processes of absorption and scattering. Exact measurements of absorption and scattering of pure water are extremely difficult to make. The actual pure absorption minima may be closer to 418 nm (Pope and Fry, 1997). However, scattering by water molecules decreases as wavelength increases (Smith and Baker, 1981), which leads to a transparency minima near 430 nm. Most phytoplankton have evolved to efficiently utilize this region of the spectrum to maximize their photosynthetic activities (Kirk, 1994; Falkowski and Raven, 1997). In the presence of sufficient light and macro- (e.g. nitrogen and phosphorous) and micro-nutrients (e.g. iron), phytoplankton growth can lead to increases in total autotrophic biomass and organic degradational products. As the total organic load increases, the amount of absorptive and scattering material increases, reducing the total photon density as well as altering the spectral nature of that density, i.e. the color of the water shifts from the blue towards the red and the water clarity is reduced. The shift in hue as a function of water column biomass has been one of the more useful relationships that have been exploited for remote sensing purposes. By examining the shift in relative terms, i.e. dividing the upwelling light from the blue region by the upwelling light in the green region, a quantitative empirical relationship between 'color' and phytoplankton biomass was found in open ocean waters (Gordon et al., 1983; Gordon, 1987; Gordon et al., 1988; Mueller and Austin, 1992). These types of relationships have been used with the CZCS data to produce the first large-scale synoptic estimates of phytoplankton biomass. This type of relationship continues to be used today with the more recent ocean color sensor (Plate 5), Sea Wide Field-of-view Sensor [SeaWiFSI. The absorption of light by phytoplankton results primarily from the light-harvesting pigments within the thylakoid membrane, as well as photoprotective pigments found in the chloroplast envelope. Chlorophyll a is the ubiquitous pigment found in all marine algae (Rowan, 1989), and as such has been used as a proxy for total phytoplankton biomass. The use of this pigment as a proxy for autotrophic biomass has been criticized because of the extreme variances in the ratio of chlorophyll a per cell (Buck et al., 1996; Stramski et al., 1999). However, the techniques for measuring chlorophyll a are relatively simple (Yentsch and Menzel, 1963; Holm-Hansen and Riemann, 1978; Bissett et al., 1997) and there are numerous empirical relationships between total chlorophyll a and phytoplankton standing stock, as well as total primary productivity. Thus, this pigment has been used for decades as the measure of phytoplankton biomass. Usage of a pigment as an indicator of biomass was also heuristically appealing to ocean color scientists because of the direct link between pigments and absorption of light in the water column. 520
This chapter will describe the basics of ocean color remote sensing. It will include a description of h o w to obtain a n d use SeaWiFS data within NASA's freely available ocean color remote sensing software. In addition, we will describe s o m e differences in m e t h o d o l o g y and touch u p o n some of the m o r e recent d e v e l o p m e n t s in the optical remote sensing field.
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PRINCIPLE
Geometrical radiometry O u r discussion starts -~with a short review of radiornetry, geometry, and radiative transfer theory. Optical remote sensing is concerned with the m e a s u r e m e n t of radiant energy (light) after a target or m e d i u m of interest has modified it. Light is defined in terms of energy units of joules (J = 1 kg m ~s ), or p o w e r units of watts (W = 1 J s ~). Alternatively, we could speak of light as individual packets called p h o t o n s or quanta (wave-particle duality is a cornerstone of m o d e r n physics (Mobley, 1994)). An einstein is equal to a mol of p h o t o n s (1 einst - 6.023 x 10 > photons; a m o r e recently accepted n o m e n c l a t u r e is 1 mol quanta - 1 einst). These definitions of light are related by the w a v e l e n g t h , the speed of light, and Planck's constant:
q-
]IC ;t
(26.1)
w h e r e q is equal to the energy of a photon; h is Planck's constant -- 6.626 x 10 ~ J s; c is the s p e e d of light = 2.998 x 10 ~ m s '; and )~ is the w a v e l e n g t h (in meters; note that c is given in m s '; usage of this formula requires that w a v e l e n g t h and s p e e d of light h a v e the correct units) of interest. The m o s t useful m e a s u r e m e n t s of light for remote-sensing p u r p o s e s are radiance (L) and irradiance (E). Radiance is operationally defined as: L
_
AQ At AA A~ A,~
(J s 1 m : sr J n m l)
(26.2)
which states that radiance is the a m o u n t of energy AQ, received in a time interval At, b y a detector of area AA, which is viewing a solid angle A~, and w h o s e w a v e l e n g t h filter passes a w a v e l e n g t h b a n d of size A)v. The m e a s u r e m e n t of a solid angle is given in steradians (sr). [t refers to the area of a sphere s u b t e n d e d by a set of radi from the sphere's center divided by the radius of the sphere squared. The best w a y to visualize the concept of a solid angle is to imagine yourself inside a sphere, at its center, holding an e m p t y p a p e r towel tube to y o u r eye. Your eye can see an area on the surface of the sphere, through the tube, of size AREA. The distance from the center of the sphere to the surface of the sphere is the RADIUS, thus the solid angle ~ = A R E A / R A D I U S : in steradians. As the total area of the sphere is 4rr(radius) 2, the solid angle of an entire sphere is equal to 47r(sr).
521
By this analogy, the remote-sensing instrument is essentially a collecting tube (the e m p t y paper towel tube of the above example) with a detector at its base (your eye). The inside of the tube is painted black to minimize photons coming from outside the desired solid angle from bouncing off the inside sides of the tube into the detector. A diffuser is typically placed before the detector, so that the detector only has to sample a fraction of the area of the diffuser to determine the total incoming radiance. The surface area of the diffusing plate has an area, A, associated with it, such that all of the terms of Equation (2) are now defined. Figure 26.1 gives a schematic drawing of such an instrument.
light detector~be
~/~,,,~
~i(~)
"~A~
filterA~ Figure 26.1. Schematic design of an instrument for measuring unpolarized spectral radiance (redrawn from MoNey (1994)).
As we are talking about the pointing of collecting tubes, we need to u n d e r s t a n d a couple of terms about directionality. A sensor looking straight d o w n is said to have a viewing angle, or nadir angle, of 0 °. As the sensor moves 'off' nadir, this angle changes in a positive direction, such that a horizontal view w o u l d be 90 °, and a vertically (upward) looking sensor would have a nadir angle of 180 °. As the sensor moves off vertical viewing, it acquires an azimuthal viewing angle, 0, which is typically measured clockwise from the instrument's (satellite) direction of travel. Ocean color sensors are called passive sensors, which means they do not have an illuminating, or active, source of light, but rather, passively collect the light coming from the planet. In order to quantify the information derived from light impacting a passive detector flying high above the earth, we first need to k n o w what the total irradiance at the area of interest on the surface of the planet is. Thus, the other useful measurement for remote sensing is downwelling irradiance. If we remove the tube from our above instrument, and set it on the g r o u n d facing upward, it would collect light from all d o w n w a r d directions. The integration of all d o w n w a r d traveling photons over all nadir and azimuthal angles is called d o w n 522
welling irradiance, E~. The detector, however, does not see the radiance equally over all solid angles. Consider a laser looking straight d o w n onto the detector (nadir angle, 0, of 0 degrees), whose beam exactly fits onto the detector. N o w consider the same laser at a 45 ° angle to the detector. The beam will be spread out u p o n the ground, and the detector sees only a fraction of the total off-nadir light as it is dispersed over a larger projected area on the ground. The dispersal of the p h o t o n density is proportional to AA cos 0. (The quantity AA cos 0 is the area that the detector projects into the plane perpendicular to the beam direction). Thus, downwelling irradiance is simply the integral of radiance over all nadir and azimuthal angles multiplied by the cosine of the nadir angle.
Why is the sky blue? We continue our discussion with the spectrum of sunlight and the impacts of a fluid m e d i u m (the atmosphere) on the downwelling light field. The visible solar irradiance at the top of the atmosphere is blue-rich (peaking in m a g n i t u d e at -450 nm). This irradiance from the sun is reduced as it passes through the atmosphere, and blue light is preferentially removed relative to red light in a clear atmosphere. The relative impact of the blue reduction becomes greater the more atmosphere the solar irradiance has to penetrate. This should be intuitively obvious for those who have seen the sun at noon and the sun at sunset. At noon the sun is directly overhead and the distance through the atmosphere is minimized, and it appears nearly white. At sunset, photons must pass through a greater v o l u m e of the atmosphere to arrive at the same point. The result is a sun that appears to be d i m m e r and shift in color towards red. The reason for the color shift and reduction in energy has to do with the inherent and apparent optical properties (lOPs and AOPs) of the atmosphere. The inherent optical properties refer to the properties of a m e d i u m that impact a photon as it travels through a finite distance of the medium. These properties do not d e p e n d on the directionality of the photons. For this discussion we are going to assume there are only three possible processes that impact a photon as it passes into a given medium. First, the material in that m e d i u m can absorb the photon, completely removing it from the incoming radiant energy. Second, the p h o t o n can be scattered by the material, changing its directionality but otherwise not impacting the radiant power. Third, it can be transmitted through the m e d i u m without interaction at all. Let us define then three processes, absorptance (A), scatterance (B), and transmittance (T) for a parallel beam of light traveling through some distance, At, of m e d i u m (Figure 26.2):
A(Z)-
•~:(,;t.) ,,(Z) '
• ~,(2)
B(Z) = ~,.(.;t) '
¢bt (2)
T(~) = ~,(~)
(26.3)
where ~,(;0 refers to the radiant p o w e r in watts (W) incident oll the medium, and ~ , qb,, and qb, refer to tile radiant p o w e r attributed to each of the processes affecting tile photons through the medium. The use of ~,
523
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U
m
td
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Figure 26.2. Geometry used to define inherent optical properties (redrawn from Mobley (1994)).
denotes the spectral dependence of each of the processes. (Note that while both absorptance and scatterance processes remove photons from the original direction of the incident beam, only absorptance truly removes the photons. Scatterance just changes the direction that they travel. However, this scattered light misses the detector placed in the path of the original beam.) We assume there are no changes in the radiant energy from internal sources of light within the medium and that no photons are absorbed and re-emitted at different wavelengths. Thus, A(;V) + BOO + TOO = 1. In our above example of the sun and atmosphere, the transmittance for blue light was less than that for green or red light. The matter in the atmosphere, i.e. oxygen, nitrogen, water vapor, clouds, dust, etc., absorb and scatter the photons traveling through it. This leads to a spectral shift in the radiant power of the total incoming light along the direct line of sight of the sun. If we are looking right at the sun, the process (absorptance or scatterance) that has the greatest impact on the reduction in the solar radiance is not completely obvious. However, if we look off angle (to the side) from the direct solar beam during a clear sky day, the dominant process removing blue light becomes obvious. As the sky is blue, we can infer that there must be some process that is preferentially removing blue light from the direct beam (scatterance), but is not completely removing the photons (absorptance). This process is called molecular scattering (often called Rayleigh scattering) and has a very strong wavelength dependence 0v 4). The inherent optical properties of absorption and scattering, a(~.) and b(~,), respectively, are defined as the absorptance and scatterance per unit distance of medium, and are given in units of m ~. Beam attenuation, c0v), is equal to the sum of a0v) and b()v). The third inherent optical property that is important is the volume scattering function, ~(~y, ;%),and refers to both the change in directionality and reduction in incident radiant power through the solid angle k~-~in Figure 26.2. Here, ~ refers to the angle that the photon travels after being scattered by the medium. ~yvaries between 524
0 ° (no change in direction) and 180 ° (complete back scattering). Integration of the v o l u m e scattering function between angle 90 ° and 180 ° yields another important quantity called total backscattering coefficient, bL~(X),which has units of m '. As we mentioned above, the satellite sensor is a passive instrument. We n o w have the terminology to more rigorously describe what the sensor is detecting. An ocean color sensor measures the upwelling radiance that is derived from the incident solar irradiance which is backscattered in the field of view of our sensor".
Biological considerations As stated above, p h y t o p l a n k t o n have adapted their photosynthetic machinery to harvest light in the blue relative to the red. Figure 26.3 shows the absorption spectra for some major bloom-forming phytoplankton found in today's oceans. The spectra were measured for phytoplankton cultures in the laboratory but illustrate the variability in p h y t o p l a n k t o n absorption due to differences in accessory pigments. Intuitively, the greater the p h y t o p l a n k t o n concentration, the lower the total light available. In a purely absorbing m e d i u m the light is r e m o v e d exponentially as a function of its absorption coefficient and the distance the photon has to travel, i.e.:
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525
where L(0) is the radiance at a boundary point; L(z) is the radiance at a distance z in a direct line from the b o u n d a r y point; a is the absorption coefficient in units m '; and Z is the distance along the direct path. This is known as Lambert-Beer's law. What is evident from Equation (4) and Figure 26.3 is that the differences in absorption coefficients will manifest themselves exponentially in the water column. In other words, the preferential removal of blue light happens exponentially as phytoplankton concentration increases. Note the m i n i m u m in the absorption spectra in the area from -520 to 600 nm. With the exception of cyanobacteria, most phytoplankton species do not have pigment complements that strongly absorb light at these wavelengths. The net color effect of increasing the phytoplankton concentration is that the water will become increasingly green. H o w green is green? And can a quantitative measure of 'greenness' be translated into an estimation of chlorophyll a a n d / o r other biological material? Using a rigorous radiative transfer code (HYDROLIGHT 4.0, http://www.sequoiasci.com/hydrolight.html) with a model of the water column IOPs as a function of chlorophyll a concentrations in typical oceanic waters (Gordon and Morel, 1983; Morel, 1991), we computed the water-leaving radiance spectra, L,,(k), as the chlorophyll a concentration increased from 0.10 to 10.0 mg Chl a m ~(Figure 26.4(a)). Note the striking 'hinge point' near 490 nm. By taking the ratio of L,,.(490) wavelengths to one of the green L,,0v) on the right of the hinge, one could imagine that a non-linear relationship could be used to map the ratio of upwelling radiance to chlorophyll a concentrations. This was the type of relationship used by the original Nimbus Experiment Team to formulate the empirical algorithm for the CZCS. The SeaWiFS algorithm (O'Reilly et al., 1998) follows the same format and is*:
chl = -0.040 + 10 l°341 3°°lX+2811x2-2°41x~ ]
(26.5)
where X = log 10 [ Rr~(490) / Rr~(555)] The SeaWiFS algorithm is a modification of the original CZCS-type algorithm as it uses remote-sensing reflectance, R,.(490) and R,.~(555), rather than normalized L,,0v), in the empirical estimation of chlorophyll a concentration (Gordon and Clark, 1981; Mueller and Austin, 1992). RT~is defined as L,,.(~,)/E~(X).Normalizing by the downwelling light field, either by the Gordon and Clark (1981) method or by division by E,0v), removes the spectral variation and directionality of the source light from the upwelling radiance, i.e. relates all measurements 'to those that would be measured were the sun at the zenith, at the mean Earth-sun distance and with the effects of the atmosphere removed' (Mueller and Austin, 1992). Figure 26.4(b) shows the R,,(X) curves from the same HYDROLIGHT runs. However, the process of absorption removes photons from the water. A satellite sensor does not view absorbed photons, rather it 'sees' the effects 526
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Figure 26.4. M o d e l e d w a t e r - l e a v i n g radiance, L,,0Q at high and low chlorophyll a concentrations. The w a t e r c o l u m n IOPs were created with a bio-optical m o d e l for Case 1 waters (Gordon and Morel, 1983; Morel, 1991) as a chlorophyll a concentration. N o t e that the Case 1 m o d e l does not include n o n - c o v a r y i n g optical constituents, i.e. river C D O M or s u s p e n d e d s e d i m e n t s f o u n d in Case 2 waters. Near-shore Case 2 waters m a y h a v e significantly different L,, spectra for the s a m e chlorophyll a concentrations. The H Y D R O L I G H T runs used the Case 1 water IOP model, with the sun at 30 ° nadir angle in a clear sky, w i n d of 5 m s ', in infinitely d e e p water, R a m a n scattering and chlorophyll a fluorescence (see the peaks around 680 nm) were included in the runs. The w i d t h of the SeaWiFS data bands are s h o w n as bars at the b o t t o m of the figure.
527
of absorption on the backscattered photons leaving the water. The decrease in L,, (Figure 26.4) in the blue is remarkably similar to the increased blue absorption of phytoplankton (Figure 26.3). In fact, the AOP of R,~ (as well as the radiometric quantity of L,,) appears to be proportional to the lOP ratio of bt7/a (Morel and Prieur, 1977). The relationship established by Morel and Prieur (1977) links an IOP (absorption) to an AOP (remote sensing reflectance). One may expect a spectral dependence of bb. Fortunately, the spectra dependence of b, is less influenced by phytoplankton than the absorption coefficient because most of the backscattering comes from very small sub-micron size particles and water itself. Molecular scattering is nearly isotropic (equal in all directions), such that water molecules have a spectrally invariant backscattering ratio of -50%. Viruses have a backscattering ratio of -20-30%, increasing slightly in the red (Stramski and Mobley, 1997). As a general rule, the greater the size of the particle beyond the molecular size, the greater the scattering, but the lower the backscattering ratio. The size of the phytoplankton load does impact the total scattering coefficient, b(;~), but the volume scattering function of phytoplankton is very weak in the backwards direction. The fraction of photons scattered backwards by phytoplankton ranges from about -0.01 to 0.20% depending on the size and wavelength (increased scattering in the red). Over a range of typical phytoplankton concentrations (away from river plumes or areas of active sediment re-suspension) the variability of b,(X)/a(;~) is mainly a function of the variability of a(X). The ratio of Lw(490)/L,,(555) is thus nearly proportional to a(555)/a(490) (or inversely proportional to a(490)/a(555)). This relationship between the ratio of absorption (an lOP) and remote sensing reflectance (an AOP) is the basis for the current satellite algorithms to estimate chlorophyll a concentration. There are many nuances to this relationship (the above relationship established by Morel and Prieur (1977) assumes that all the optical constituents co-vary with chlorophyll; this is obviously not true for vast sections of the coastal ocean), and the term 'nearly proportional' in the last sentence of the above paragraph is cause for great angst and research in the ocean color community. However, we have addressed the basics of ocean color remote sensing and will now focus the remainder of this chapter on the tools necessary to acquire and use satellite data from NASA. These tools will be demonstrated with actual images, and compared against in situ data so we can briefly discuss issues of validation.
4, 4, 4, 4, , 4, H A R D W A R E ,
SOFTWARE,
AN D DATA
Hardware requirements Image processing and analysis are computationally intensive processes. The tasks of navigation, atmospheric correction, re-mapping, and image manipulation typically require workstation caliber computers to accomplish. For our purposes, a workstation refers to a computer built around a
528
RISC chip with a UNIX operating system, i.e. SGI 02 or SUN UltraSPARC. While the power of PC-type computers has dramatically increased in the past decade, the memory requirements and execution speed have yet to match those of the workstations. However, this is a rapidly changing field and the latest PC-type machines with the LINUX operating systems may yet prove to be sufficient for image processing'. NASA currently makes available a free software package to process SeaWiFS images as part of the Mission to Planet Earth [MTPE] program. This package is called SeaWiFS Data Analysis Software [SeaDAS], and it can be downloaded from http://seadas.gsfc.nasa.gov. While there are other image processing packages available, our discussion will focus on SeaDAS, as it is free and can be used with a currently operational satellite sensor (SeaWiFS). There are many other ocean color satellites being planned (and one that has just been launched, i.e. MODIS), but the data streams are not currently available. The following discussions may become dated soon after publication, as the tools, techniques, and equipment are constantly changing. Our discussion should be viewed in the context of the process required to acquire an image from a public (NASA) image database, and used to understand some of the basics of retrieving and remapping satellite data to usable images. The reader is referred to a more complete text on remote sensing and image processing (Schowengerdt, 1997) for further information on techniques and algorithms. Oil the NASA web site, one will find the suggested requirements for computational equipment. These will change as SeaDAS changes and the computational abilities of hardware and software change. While you may choose the minimum system requirements that NASA suggests, as a general rule, more is better than less in image processing. At the very least opt for more RAM and disk memory than the minimum requirements. The reason for the increase in memory is that when analyzing a time series of satellite images you may load and display multiple images at once, which will rapidly take up RAM. If you do not have sufficient RAM, most computers are set up to use disk memory as virtual RAM (also called SWAP memory). SeaDAS is written in Interactive Data Language [IDL], and our experience with this software suggests that is does not handle swapping very efficiently. The net result is that your system may freeze up, or your process may completely blow up, causing a loss of time and data, as well as endless frustration. The increase in disk space results from the fact that a single Level 1A" image and its Level 2r processed products may be as large as 250 MB. This is without creating any data products using other algorithms, or creating publishable images.
Software requirements The required s o f t w a r e to run S e a D A S are: • •
Operating systems: SGI: IRIX 6.3 or 6.5, SUN: Solaris 2.6 or 2.7, or kinux Required software: IDL S.I or 5.2 (from Research Systems, Inc.; http://www.rsinc.com)
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• Software libraries: HDF 4.0r2 (included in SeaDAS) SeaDAS does not require the full version of IDL and can be compiled solely with the runtime version of IDL. This may save some money on the initial start up of using SeaDAS. The disadvantage to using the runtime version is that you lose all of the functionality of IDL, which does have some powerful analytical tools.
Data acquisition SeaWiFS data can be acquired from NASA's Distributed Active Archive Center [DAAC], and can be ordered online by following the ocean color links at http://daac.gsfc.nasa.gov/. SeaWiFS is a commercial instrument flying on Orbimage's (http://www.orbimage.com/) Orbview-2 spacecraft. NASA purchased data rights for its researchers prior to the satellite's launch. However, there are restrictions on how the data may be used. As long as you are doing non-profit research it is quite easy to become an Authorized SeaWiFS Data User. The links for the required documentation to become an Authorized Data User are on the Ocean Color page at the DAAC web site. Once you become an Authorized User, use the SeaWiFS data page (http://daac.gsfc.nasa.gov/data/dataset/SEAWIFS/index. html) to browse for the images you wish to acquire. New users must register here as well. Once you have registered with the DAAC, you are ready to acquire your images.
APPLICATION Natural dynamics in microbial communities reflect biological responses to environmental fluctuations (variations to light, temperature, shear, and nutrients), trophic interactions, and physical transport processes such as turbulent mixing and advection. This has made characterizing the ecology of natural microbial communities difficult. Remote sensing provides a tool that can provide information over time/space scales not possible using traditional sampling approaches from ships (Plate 6). This has fundamentally changed our view of microbial dynamics of the oceans and provides the foundation for adaptive sampling of biological communities in the future (Schofield et al., 1999). Despite much promise, scientists should cautiously view the information provided by satellite maps. We are going to demonstrate the power and pitfalls of ocean color data with an example from an active research program in the New York Bight [NYB]. Data were collected by the Coastal Ocean Observation Laboratory [COOL] at the Long-term Ecosystem Observatory [LEO-15], which is located off the central coast of New Jersey. The LEO-15 system is a coupled ocean observation/modeling system being constructed to acquire long-term high-resolution measurements from marine to coastal habitats (http://marine.rutgers.edu/cool). Currently the LEO-15
530
observation network consists of satellites, aircraft, radar, meteorological sensors, subsurface observation nodes, moorings, research vessels and a u t o n o m o u s u n d e r w a t e r vehicles. The system collects data from the Mullica R i v e r / G r e a t Bay Estuarine Reserve and across the N e w York Bight. The data described here were collected as part of a study focused on s u m m e r upwelling. We will n o w walk through the process of obtaining an image from the DAAC, processing the image, and then briefly analyzing the data in context of an ongoing coastal oceanographic program.
Obtaining an image ~ We are going to start by trying to obtain an image of tile East Coast of the United States on July 16, 1999. Go to the DAAC web site (http://daac.gsfc.nasa.gov/data/dataset/SEAWIFS/index.html) and follow the links: ---> Data Products --+ LAC (local area coverage, 1 km resolution) -9 L1A HRPT (Level 1A data from the High Resolution Picture Transmission IHRPT] stations) -9 HNSG (NASA G o d d a r d Space Flight Center, Greenbelt, Maryland, USA HRPT station) --+ 1999 (data from the year 1999) -9 July ---> S1999197171620.L1A HNSG By clicking on this link, y o u will be s h o w n a browse image of the HRPT data collected by the receiving station as the satellite was passing over God dard. The browse image is an un-navigated pseudo-color image, which allows the viewer to see if the site of interest is in the scene and visible through the clouds. The image file name is from the time stamp on the !mage, i.e. day of year 197 (July 16th) of year 1999. Order the image by following the links at the top of the page, and be sure to request all of the meteorological data in the process. The best way to receive the data is via FTP. When the data are ready you will receive an email with instructions on how to retrieve the data from the DAAC FTP server. The data will be available in compressed form, which you must uncompress on y o u r computer after d o w n l o a d i n g it.
Processing the image In the directory where y o u have u n c o m p r e s s e d the image on your computer, start SeaDAS. This will place the SeaDAS Main Menu GUI on y o u r desktop. On this interface choose: -~
Process -9 SeaWiFS --> 12gen (L2 file generation)
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On the L2 Products GUI you want to select the file that you ordered and uncompressed (S1999197171620.LIA HNSG), and give a name for the output file, e.g. S1999197171620.L2. You can use all the parameter defaults. We would suggest using the meteorological and calibration files that came with the image in the MET file parameter block. Select the Run Button. Once the image has finished processing, select the Quit Button and return to the SeaDAS Main Menu GUI interface. Now choose: -9 Display -9 seadisp (General image and graphics display) This starts the Seadisp Main Menu GUI. Select: -9 Load -9 SeaWiFS which starts the Product Selection GUI. Select the file that you created with the 12gen routine, and afterwards select the chlor_a check mark under the Products sub-page. This brings up the Band Selection GUI. You can display this product, however, it will not be mapped into a projection that is easy to use. Instead, on the SeaDisp Main Menu GUI select: -9 Functions -9 Projection which starts the SeaDisp Projection GUI. Select the chlor_a band, and drop down to the Projections button and choose Cylindrical. Below this input panel, set the north and south latitude and east and west longitude coordinates for the desired limits of the image (Plate 6 limits are approximately 30.20 ° N and 30.75°N, by 74.50°W and 73.75°W), and click on the Go Button. A new band will be displayed in the Band List Selection GUI called Mapped -chloro a (your filename). Display this image. A new window will be displayed with the mapped data. The Function Button will allow you to add coastlines, color palettes, output the data, etc.
Basic image analysis The largest (non-seasonal) variations in ocean temperatures along the New Jersey coast are caused by episodic summertime upwelling events forced by southwesterly winds associated with the Bermuda High. Off the southern coast of New Jersey, topographic variations associated with ancient river deltas direct the upwelled water to evolve into an alongshore line of three recurrent upwelling centers that are co-located with historical regions of low dissolved oxygen [DO]. Remote sensing has been a key tool in mapping the cold, nutrient-rich, upwelled water. This nutrient-laden water supports large phytoplankton blooms when exposed to sunlight, which in turn provides a steady flux of organic material to the underlying bottom waters. Under the right conditions, the supply of organic material exceeds the supply of oxygenated waters, with subsequent remineralization leading to low oxygen conditions. The coherence between the cold 532
upwelling water, as depicted by the sea surface temperature [SST] m i n i m u m (Plate 6(b)) near N o d e A, and the increase in SeaWiFS estimated chlorophyll a concentration (Plate 6(a)) is clearly evident. This relationship between cold nutrient-rich water and sea surface pigments (when there is sufficient sunlight for autotrophic growth) in the NYB is an extension of the more general relationship, also depicted in Plate 5, over m u c h of the world's oceans. Use of remote sensing data places the LEO-15 field p r o g r a m in the context of the larger oceanic environment, providing the necessary information with which to view the in situ data (Plate 7). The in situ data of absorption and scattering collected along the transect line running to the north of N o d e A (Plate 6) confirms our theoretical interpretation of the effect of increasing biomass on the relative changes in ratios of L,,(;~), a(X), and bb(Z). As mentioned above (and seen in Equation (5)), we expect the ratio of L,,(490)/Lw(555) to increase (seen as increasing chlorophyll a in the SeaWiFS image) as the ratio of a(490)/a(555) decreases, with very little spectral change in b~(X) (Figure 26.5; note that the difference in wavelength for b,(;t,) is the result of the different available bands on the in situ instruments). Plate 7 also demonstrates a source of error in the satellite-derived chlorophyll a (and SST as well). The ocean is not a two-dimensional surface, but rather a three-dimensional fluid medium. However, satellites can only 'see' some small distance into the surface of the ocean. The depth at which information can be retrieved from remotely sensed data d e p e n d s on the penetration depth of the light, which in turn is a function of the water column IOPs. Clearly, some wavelengths are going to penetrate deeper than others and the d e p t h of penetration d e p e n d s on h o w m u c h 'stuff' is in the water. In h o m o g e n e o u s water, there is an exponential decay ~' in the p h o t o n density as light penetrates d o w n w a r d . Photons are backscattered into the u p w a r d direction at each depth, and there is also an
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exponential decay in the photons traveling back toward the surface. This two-way travel by photons means that the information derived by the spectral change in water-leaving radiance is an integration of the water column's IOPs, heavily weighted by the near-surface values. Thus, any empirical approach relating in situ optical constituents to water-leaving radiance becomes an integrated estimate over the distance that the photons have penetrated, i.e. the chlorophyll a estimates in Plate 6(a) are integrated near-surface values. The vertical d e p e n d e n c e of the lOPs in Plate 7 is lost in the satellite data. But h o w deep does the satellite 'surface' water extend? Based on the estimate of chlorophyll a, one could estimate the attenuation of d o w n welling irradiance, K, per unit distance and use an equation similar to Equation (4) to estimate the penetration depth of the light. In general, -90% of the light that leaves the surface waters come from the first diffuse attenuation depth (where diffuse attenuation depth is defined as 1/K). Note that different colors of light integrate over different distances in the water because the diffuse attenuation K depends strongly on wavelength. The additional complications owing to vertical variations of the IOPs have m a d e it necessary to use empirical relationships like Equation (5) to estimate chlorophyll. This formulation will have wide error bars, but for the time being is a reasonable, all-purpose, algorithm for most open ocean conditions. The algorithm in Equation (5) was primarily developed for open ocean conditions where the color signal is solely a function of the in situ produced organic material. In addition, it assumes that all of the organic colored constituents, i.e. phytoplankton, dead phytoplankton, phytodetritus, Colored Dissolved Organic Matter [CDOM], co-vary with each other. These kinds of waters are often referred to as Case 1 waters (Morel and Prieur, 1977), and they represent a majority of the world oceans (N80 to 90%). Coastal waters have additional sources of color that do not necessarily co-vary with primary production. The additional color constituents include re-suspended sediments, bottom reflection, river-derived CDOM, etc. Coastal waters that have non-covarying optical constituents are often referred to as Case 2 waters. For these more complicated coastal ocean conditions, new algorithms that specifically address the vertical structure of water column IOPs are being developed (Gould and Arnone, 1998). These usually require some additional information on in situ IOPs at the time of the image collection, which are subsequently used in conjunction with reflectance maps to derive three-dimensional IOP estimates. in Plate 6, if we were to imagine a transect parallel to the N line through N o d e A, we would see chlorophyll a decreasing with increasing temperature. This conforms to our general interpretation of organic material and water temperature (Plate 5). Northeast of N o d e A along the N transect line is a different story. The cold water seems to split an area of w a r m e r water, such that transecting from the coast to deeper water yields a w a r m --~ cold --~ warm line. We would expect to see a commensurate low chlorophyll high chlorophyll --+ low chlorophyll SeaWiFS plot. However, we see a much higher a m o u n t of estimated chlorophyll in water near the coast at temperatures near 20°C than we do offshore at similar temperatures. This 534
higher chlorophyll estimate nearer to the shore results from optical constituents that do not co-vary with the chlorophyll a concentration, i.e. reflectance of light off the bottom, higher concentrations of CDOM coming from the rivers and estuaries, and re-suspended sediments from the bottom. In Case 2 waters, great care must be taken in divining detailed information from simple algorithms and SeaWiFS data. Table 26.1 shows actual chlorophyll a measurements along the N line in Plate 6. Notice the increasing error in the SeaWiFS estimate as we move from offshore to onshore. The errors in the SeaWiFS estimates appear to stabilize around ~ 5 m g chl a m ~ at approximately 10 km offshore. SeaWiFS estimated chlorophyll a concentrations greater than this should be analyzed carefully in these types of waters.
Table 26.1 Comparison between satellite-derived chlorophyll a and in situ HPLC surface measurements of chlorophyll a. Sample corresponds to stations in Figure 26.5. The decreasing difference between in situ HPLC chlorophyll and SeaWiFS estimates results from the decreasing influence of non-covarying optical constituents, i.e. sediments, estuarine CDOM, etc., on the upwelling radiance signal. Distance from shore (km)
SeaWiFS chl a (mg m 3)
HPLC chl a (mg m 3)
3.5 6.5 10.0
17.92 8.86 4.04
8.26 6.80 4.48
FUTURE DIRECTIONS There are m a n y research efforts trying to develop more accurate lOP algorithms from ocean color data to address Case 2 water problems, as well as to derive the concentrations of other optically active constituents, e.g. CDOM. NASA's Sensor lntercomparison and Merger for Biological and Interdisciplinary Oceanic Studies (SIMBIOS; h t t p : / / s i m bios.gsfc.nasa.gov/) p r o g r a m is one of the mechanisms by which a large fraction of this work is funded. Their web site is a good starting point for the latest information on ocean color algorithm development. There are other ocean color sensors, besides the SeaWiFS sensor, currently operating. These included the NASA Moderate Resolution hnaging Spectroradiometer (MODIS), the Indian Remote Sensing Satellite Modular Optoelectronic Scanner (MOS) and Ocean Color Monitor (OCM), the European POLarization and Directionality of the Earth's Reflectance (POLDER), and the Taiwanese Ocean Color imager (OCI). The acquisition and manipulation of these data streams are a bit more difficult. However, more intensive studies into ocean color may find the usage of multiple remote sensing data streams a means to acquire more complete temporal and spatial coverage of a studv site, as well as providing crosscalibration of the data streams. 535
CONCLUSIONS We h a v e r e v i e w e d the basic tenets of o c e a n color r e m o t e sensing. By e x p l o r i n g h o w light penetrates the w a t e r c o l u m n a n d h o w the optical constituents i m p a c t the light as it travels t h r o u g h the water, w e h o p e to p r o v i d e the basic u n d e r s t a n d i n g of the v a l u e a n d limitations of o c e a n color data. While w e h a v e s h o w n w h e r e to obtain SeaWiFS images, a n d s o f t w a r e to process these images, the r e a d e r s h o u l d use this c h a p t e r as just one of m a n y references on ocean color r e m o t e sensing. There is a large scientific difference b e t w e e n m a k i n g color pictures a n d u n d e r s t a n d i n g the d r i v i n g processes b e h i n d spatial variations of w a t e r - l e a v i n g radiance. R e m o t e sensing is an i n d i s p e n s a b l e tool in o c e a n o g r a p h y , a n d u s e d correctly can greatly facilitative the interpretation of in situ and l a b o r a t o r y data.
Acknowledgement This w o r k w a s s u p p o r t e d b y the Office of N a v a l Research.
Endnotes ' Gross primary production minus plant respiration. 2 While a complete description of radiometry and hydrological optics is beyond the scope of this chapter, there are some basic tenets of ocean optics that must be covered in order to proceed with the utilization of remote sensing data. For a more complete discussion of hydrological optics and its impacts on photosynthesis, see Mobley (1994) and Kirk (1994). Remember we have assumed that there are no internal sources of light or absorption/emission processes, e.g. solar stimulated fluorescence. The operational coefficients of this algorithm have changed. It should be noted that these coefficients frequently change, depending on new processing, atmospheric correction, etc., as well as region and seasonal changes for site- and timespecific studies. This is relative of course. Digital image processing has been a recognized field of endeavor for the last several decades, at the beginning of which supercomputers were but a fraction of the power of today's PCs. However, as the power of computers has increased, so have the demands of image processing, i.e. greater image size and resolution, more wavelength bands of information, etc. " Level 1A [L1A]: reconstructed, unprocessed instrument data at full resolution, including radiometric and geometric calibration coefficients and georeferencing parameters (i.e. platform ephemeris) computed and appended, but not applied to the L0 data (see http://seadas.gsfc.nasa.gov/doc/sds faq.html#G levels). Level [L2]: derived environmental variables at the same resolution and location as the L1 data. The next two sections are valid as of May 1, 2000. Future updates to the DAAC and SeaDAS software may render them obsolete. However, the processes of obtaining an image and deriving products will be similar, so the following descriptions may be used as a reference. " Equation (4) with an attenuation term, K, instead of absorption, a. K is a slight modification of a resulting from the effects of backscattering and the average direction of the photon density.
536
References Acker, J. G. (1994). Volume 21, The Heritage of SeaWiFS: A Retrospective of the CZCS NIMBUS Experiment Team (NET) Program. NASA Center for AeroSpace information: Linthicum Heights, MD. Bissett, W. P., Patch, J. S., Carder, K. L. and Lee, Z. (1997). Pigment packaging and chlorophyll a-specific absorption in high-light oceanic waters. Limnol. Oceanogr. 42, 961-968. Buck, K. R., Chavez, E P. and Campbell, L. (1996). Basin-wide distributions of living carbon components and the inverted trophic pyramid of the central gyre of the North Atlantic Ocean, summer 1993. Aquatic Microbial Ecol. 10, 283-298. Denman, K., Hofmann, E., Marchant, H., Abbott, M. R., Bates, T. S., Calvert, S. E., Fasham, M. J., Jahnke, R., Kempe, S., Lara, R. J., Law, C. S., Liss, E S., Michaels, A. E, Pedersen, T. E, Peha, M. A., Platt, T., Sharp, J., Thomas, D. N., Scoy, K. A. V., Walsh, J. J. and Watson, A. J. (1996). Marine biotic responses to environmental change and feedbacks to climate. In: Climate Change 1995: The Science of Climate Change (J. T. Houghton, L. G. M. Filho, B. A. Callander, N. Harris, A. Kattenberg and K. MaskelI, Eds), pp. 485-516. Cambridge, University Press, Cambridge. Falkowski, P. G. and Raven, J. A. (1997). Aquatic Photosynthesis. Blackwell Sciences, Inc., Malden, MA. Gordon, H. R. (1987). Bio-optical model describing the distribution of irradiance at the sea surface resulting from a point source embedded in the ocean. Appl. Opt. 26, 4133-4148. Gordon, H. R. and Clark, D. K. (1981). Clear water radiances for atmospheric correction of coastal zone color scanner imagery. Appl. Opt. 20, 4175-4180. Gordon, H. R. and Morel, A. (1983). Remote Assessment of Ocean Color for htterp~vtation of Satellite Visible Imagery, a Review. Springer-Verlag, New York. Gordon, H. R., Brown, J. W., Brown, O. B., Evans, R. H. and Clark, D. K. (1983). Nimbus 7 CZCS: reduction of its radiometric sensitivity with time. Appl. Opt. 22, 3929-3931. Gordon, H. R., Brown, O. B., Evans, R. H., Brown, J. W., Smith, R. C., Baker, K. S. and Clark, D. K. (1988). A semianalyfic radiance model of ocean color. J. Geophys. Res. 93, 10,909-10,924. Gould, R. W. and Arnone, R. A. (1998). Three-dimensional modelling of inherent optical properties in a coastal environment: coupling ocean colour imagery and in situ measurements, hit. J. Remote Seusing 19, 2141 2159. Holm-Hansen, O. and Riemann, B. (1978). Chlorophyll a determination: improvements in methodology. OIKOS 30, 438-447. Kirk, J. T. O. (1994). Li[~ht and Photosynthesis itl Aquatic Ecosystems. Cambridge University Press, Cambridge. Moblex; C. D. (1994). Lixq# amt Water. Academic Press, San Diego, CA. Morel, A. (1991). Light and marine photosynthesis: a spectral model with geochemical and climatological implications. Prog. Oceanogr. 26, 263 306. Morel, A. and Prieur, L. (1977). Analysis of variations in ocean color. Limnol. Oceanogr. 22, 709 722. Mueller, J. L. and Austin, R. W. (1992). Volume 5, Ocean Optics Protocols for SeaWiFS Validation. NASA Center for AeroSpace Information, Greenbelt, MD. O'Reilly, J. E., Maritorena, S., Mitchell, B. G., et al. (1998). Ocean color chlorophyll algorithms for SeaWiFs. ]. Geophys. Res., 103, 24, 937-24, 953. Parsons, T. R., Takahashi, M. and lqargrave, B. (1984). Biola%ical Oceanognq#Hc Processes. Pergamon Press, New York. Pope, R. M. and Fry, E. S. (1997). Absorption spectrum (380-700 nm) of pure water. 1I. Integrating cavity measurements. Appl. Opt. 36, 8710-8723.
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Rowan, K. S. (1989). Photosynthesis Pigments of Algae. Cambridge University Press, Cambridge. Schofield, O., Grzymski, J., Bissett, W. P., Kirkpatrick, G., Millie, D. E, Moline, M. and Roesler, C. S. (1999). Optical monitoring and forecasting systems for harmful algal blooms: possibility or pipe dream? J. Phycol. 35, 1477 1496. Schowengerdt, R. A. (1997). Remote Sensing; Models and Methods for Ima~c Processing. Academic Press, San Diego, CA. Smith, R. C. and Baker, K. S. (1981). Optical properties of the clearest natural waters (200-800 nm). Appl. Opt. 20, 177-184. Stramski, D. and Mobley, C. D. (1997). Effects of microbial particles on oceanic optics: a database of single-particle optical properties. Limnol. Oceano~r. 42, 538-549. Stramski, D., Reynolds, R. A., Kahru, M. and Mitchell, B. G. (1999). Estimation of particulate organic carbon in the ocean from satellite remote sensing. Science 285, 239-242. Yentsch, C. S. and Menzel, D. W. (1963). A method for the determination of phytoplankton chlorophyll and phaeophytin by fluorescence. Deep-Sea Res. 10, 221-231.
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27 Microbial Public Health Indicators in the Marine Environment D a l e W Griffin Department of Morine Sciences,University of South FloridG St Petersburg,Florido, USA
CONTENTS Introduction Total and fecal coliforms
Enterococci Clostridium per~ringens F+ specific RNA coliphage Conclusions
eeeeee
INTRODUCTION In an article published in 1885, Dr Theodore Escherich w o n d e r e d w h y so little research was directed at identifying microorganisms in h u m a n excrement and the intestine, given the a m o u n t of research directed towards identifying microorganisms that cause disease (Escherich, 1885). Dr Escherich obviously sensed a connection between the presence of microorganisms in the intestines and disease. One h u n d r e d and fifteen years later the central question in aquatic public health microbiology is, what intestinal microorganism when found in water, is an ideal indicator of risk to h u m a n health? In regard to aquatic public health microbiology, research has yet to provide an answer. While an ideal indicator has not been identified, several microbes are currently used in the United States of America, to assess biological marine water quality. These microbes include the coliforms, enterococci and Clostridium pelfringens. Water quality is assessed by determining the numbers of these organisms in a given quantity of water. This is typically expressed as the n u m b e r of colony-forming units (CFUs) per 100 ml of water. Threshold concentration levels are based on research that has demonstrated an unacceptable risk to h u m a n health when that concentration of bacteria is met or exceeded (Cabelli et al., 1983). The United States Environmental Protection Agency (USEPA) currently r e c o m m e n d s that enterococci should be used as an indicator in
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marine waters with a guidance level of 35 enterococci CFU (geometric mean) per 100 ml for five samples spaced over a 30 day period (Dufour et al., 1986). For individual samples a guidance level of 104 enterococci CFU per 100 ml of sample has been recommended. The State of Hawaii has adopted Clostridium spp. as an indicator with a guidance level < 5 CFU per 100 ml (geometric mean), based on research that has demonstrated that this organism is a better indicator (Clostridium spp. are spore-formers and can survive longer in marine environments) than the coliforms or enterococci in its tropical waters (Fujioka and Shizumura, 1985). Although it is well d o c u m e n t e d that the coliforms are not good indicators of marine water quality, particularly in the tropics, a n u m b e r of States including Florida currently utilize coliforms to assess water quality. Coliform research has demonstrated rapid die-off in marine waters (and, in contrast, long-term survival in marine waters), isolation from areas far r e m o v e d from h u m a n activity and the ability of some marine bacteria to mimic coliforms on identification media (Santiago-Mercado and Hazen, 1987; Hazen and Toranzos, 1990). Within the United States no one indicator or indicator level/limit has been adopted by all states that monitor marine water quality. Seventeen states still utilize fecal coliforms to access water quality and only seven have adopted enterococci as an indicator (USEPA, 1998). In addition m a n y States only monitor a n u m b e r of beaches utilized for recreation and some do not monitor at all (NRDC, 1998). Our research group has screened water samples throughout the State of Florida, including brackish and marine ecosystems for the mentioned indicators (in addition to a n u m b e r of bacteriophage, h u m a n protozoan and viral pathogens) (Lipp et al., 1997; Griffin et al., 1999; Lipp et al., 1999; Griffin et al., 2000). We have noted that the higher levels of total and fecal coliforms are often found in areas far removed from h u m a n activity which suggests animal input, long-term survival and transport. We have also found m a n y marine environments where total and fecal coliform levels were below the D O H standards, while h u m a n enteroviruses could be detected by molecular techniques such as reverse transcriptase polymerase chain reaction (RT-PCR, Table 27.1) and cell culture. Enterococci levels at a n u m b e r of these sites were above the USEPA single sample limit. In addition, while most sites were only sampled once or at a frequency of less than five times a month, many of these sites had numbers of enterococci above the r e c o m m e n d e d USEPA geometric mean standard. When Clostridium pet2friHgens were detected (at a lower frequency than the enterococci) it was usually at sample sites containing the highest overall numbers of indicators. These research projects indicate (currently) a trend of enterococci being a better indicator than Clostridium perfringens and Clostridium perfrin~,~ens being a better indicator than the coliforms. In summary, in a variety of marine settings around Florida total and fecal coliform numbers have indicated acceptable water quality, when a n u m b e r of sites were in violation of the USEPA enterococci r e c o m m e n d e d limits, the Hawaiian Clostridium pet~[ringens standard a n d / o r were RT-PCR positive for h u m a n pathogenic viruses. This approach to marine water quality monitoring (monitoring sites for a broad range of indicators, alternate indicators and pathogens) may be a 542
Table 27.1 Bacterial indicator and human virus single sample results from 19 different sites in the Florida Keys Florida Keys sites
Total coliforms CFU per 100 ml
A B C D E F G H l j K L M N O P Q R S
328 15 480og log 36 390 30 11 40 60 330 14 20 11 168 0 30 710 1410
Fecal coliforms CFU per 100 ml
Enterococci spp. CFU per 100 ml
56 9 770 6 19 294 26 10
78 12 800 0 1 80 30 18 51 27 240 80 0 23 94 0 {} 63 680
20
44 220 1 5 6 150 0 {} 130 6{)0
Clostridium
perflringens CFU per 100 ml
5 5 26 0 1 6 22 1 4 0 1 0 0 3 0 0 o 3 520
RT-PCR detection of human viruses. Presence/Absence 1~[ E,H E,H,N H E,H E,N E,H E,H H E E,H E,f I E E,H E E E,H E
CFU = colony forming units. tG
ovcrgrov, th vchich prevented accurate enLlnlur~tiuFi. E Enteroviruses (a group including Polio, Coxsackie A and B, Echo and Entero). 1I Hepatitis A \iruses.
N Norwalk \ iruscs. State of Florida Department ot Health, single sample total and Jecal coliform limits are 2400 CFU per 100 ml and S,00 CFU per I00 ml respectively. USEPA recommended enterococci spp. single sample limit is 104 CFU per 101) ml. The Hawaiian CIo,tridim~ spp. single sample limit is 50 CFU pet" 100 ml.
valuable tool in identifying the best water quality indicator for a given region. A l t h o u g h we do not believe that any of the current indicators are ideal for all situations, this a p p r o a c h has helped us m o v e t o w a r d a goal of d e t e r m i n i n g an accurate marine indicator s y s t e m for the state of Florida. This chapter will present several standardized m e d i a - b a s e d m e t h o d s used to detect bacteria that are currently used as indicators of marine water quality in the United States (total and fecal coliforms, enterococci and Clostridium pecfringens). Also included is an F-specific R N A coliphage assay (a g r o u p of v i r u s e s / b a c t e r i o p h a g e that utilizes the coliform Escherichia coli, which can p r o d u c e pili, as a host) that has been used by a n u m b e r of researchers to assess biological w a t e r quality (Hsu ct al., 1995; Griffin eta[., 2000). The value of the F'-specific R N A coliphage assay is that it m a y assist in identifying the source of contamination (animal versus 543
human). For those w h o are interested in working with media-based methods for the identification of indicators in water samples, a good overall reference source is Standard Methods for the Examination of Water and Wastewater (APHA, 1998).
T O T A L A N D FECAL C O L I F O R M S Coliform bacteria are a group of bacteria that are the normal microflora in the intestines of w a r m - b l o o d e d animals. Species of bacteria within the various genera (Escherichia, Klebsiella, Enterobacter, Citrobacter, Salmonella, Hafnia, etc.) cover the full range of non-pathogenic and pathogenic intestinal microflora. The bacteria Escherichia coli (a fecal coliform within the total coliform group) has long been considered the golden standard in microbial water quality monitoring. They have been used to assess drinking water quality since the late 1800s. As stated, a n u m b e r of states still use the coliforms to determine water quality in marine recreational waters. One concern of using the coliforms to assess water quality is that they rapidly die off or enter into a state where they are viable but nonculturable u p o n entering into marine waters (Garcia-Lara et al., 1991; Huanca et al., 1996). In contrast, some research projects have d e m o n strated the long-term survival of coliforms in marine waters or have identified viable coliforms in tropical marine settings such as reefs and or shallow w a r m nutrient-laden sediments (Gerba and McLeod, 1976; Santiago-Mercado and Hazen, 1987; Perez-Rosas and Hazen, 1988; Byrd and Colwell, 1993; Hagler et al., 1993; Paul et al., 1995; Griffin, 1999). Another concern that we have noted is that m a n y Vibrio spp. and Pseudomonas spp. marine isolates can mimic coliforms on coliform media, which results in false positive counts. There are two books that are recomm e n d e d as good references for coliforms and public health, the first covering the history, problems and future of coliforms in the world of drinking water assessment, is Gleeson and Gray (1997) and the second, which covers standard m e t h o d s of identification and verification, is Lisle (1993).
E q u i p m e n t and reagents Membrane filtration equipment 1. Lubricated rotary vane-type v a c u u m p u m p (Gast Manufacturing, Buckinghamshire, England) - - purchased through Fisher Scientific (Norcross, GA), product # 01-094-22. 2. Vacuum manifold - - Pall Gelman Sciences (Ann Arbor, MI), product # 4205. 3. 47 m m magnetic filter funnels - - Pall Gelman Sciences (Ann Arbor, MI), product # 4242. 4. 0.45 micron, 47 m m presterilized cellulosic filters - - Micron Separations Inc. (Westborough, MA), product # E04WG047S4.
544
60 m m x 15 m m disposable Petri dishes - - Fisher Scientific (Norcross, GA), product # 08-757-13A. 6. Miscellaneous e q u i p m e n t - - incubators (one large capacity 36.0°C for the total coliforms and the coliphage; one 41.0°C for the enterococci; one 44.5°C waterbath for the fecal coliforms and Clostridium perfringens), forceps, alcohol, alcohol b u r n e r / b u n s e n burner, p e r m a n e n t marker for plate labeling. Tubing and b e a k e r s / b e a k e r caps for v a c u u m p u m p overflow traps. Sterile H20 and sterile rinse bottle. .
Total coliform media 1. m ENDO AGAR LES (Difco Laboratories, Detroit, MI) - - purchased through Fisher Scientific (Norcross, GA), product # DF0736-17-2. 2. Ethyl Alcohol USP, Absolute - - 200 proof, AAPER Alcohol and Chemical Co. (Shelbyville, KY). 3. E.coli C - - American Type Culture Collection (Rockville, MD), product # 13706. 4. m ENDO media recipe per Difco Laboratories, Detroit, MI (1 1) • m ENDO AGAR LES 51.0 g • Ethanol 20.0 ml • Distilled or deionized H20 980.0 ml • This media is boiled to dissolve the solids and is not autoclaved. Let cool to -48.0°C and p o u r plates.
Fecal coliform media 1. m FC AGAR (Difco Laboratories, Detroit, MI) - - purchased through Fisher Scientific (Norcross, GA), product # DF0677-17-3. 2. Rosolic acid (Difco Laboratories, Detroit, MI) - - purchased through Fisher Scientific (Norcross, GA), product # DF3228-09-1. 3. Sodium hydroxide - - Fisher Scientific (Norcross, GA), product # $318-3. 4. E.coli C - - A m e r i c a n Type Culture Collection (Rockville, MD), product # 13706. 5. m FC media recipe per Difco Laboratories, Detroit, MI (1 1) • mFCAGAR 52.0g • Distilled or deionized H,O 1.0 1 • Boil to dissolve the solids then add • 1~ rosolic acid in 0.2 N N a O H 10.0 ml • Heat for 1 min, do not autoclave, cool to N48.0°C and p o u r plates.
Assays M e m b r a n e filtration
1. The m e m b r a n e filtration apparatus is assembled as in Figure 27.1. As an alternative to the v a c u u m p u m p and traps, a faucet v a c u u m and sink can be used (this is a cheaper alternative but the m a x i m u m v a c u u m generated is less than can be obtained with a p u m p , which results in longer filtration times). 545
e, 4,J i
"1"
°m
,,Q
O to °m
1¢
i
Figure 27.1. Typical membrane filtration setup using a vacuum pump and traps.
2. Sterile filtration funnels are used for each water sample to be evaluated. 3. When placing and removing filters from the filtration funnels the forceps should be sterilized by dipping in alcohol and flaming with a Bunsen burner or alcohol burner. 4. A negative control sample should be taken prior to using the filtration funnel by placing a m e m b r a n e filter in the funnel and rinsing the funnel with sterile H20. The filter is then r e m o v e d and placed cell side up on the appropriate media. Repeat this step for each media type used. 5. When evaluating various aliquots of the water sample, start with the smallest volumes and work up to the larger volumes. Sample volumes of 50.0ml and 5.0ml (adequate volumes in most marine environmental settings) are filtered through m e m b r a n e filters. All sample aliquots are assayed in duplicate. 6. If the environmental sample being evaluated is u n d e r the direct influence of fecal waste, an additional v o l u m e of 0.5 ml of sample can be a d d e d (add enough sterile H~O to cover the filter and then add the 0.5 ml of sample to ensure even distribution of the aliquot over the entire filter surface). 7. After each aliquot is filtered the filtration funnels are rinsed with sterile water, and the filters are removed from the funnels with sterile forceps and placed on appropriate media.
Total coliforms 1. Water samples are filtered as described above. 2. The filters are placed on m E N D O media and incubated for 24 h at
37°C.
546
3. The colonies that produced a metallic sheen are counted as total coliforms (APHA, 1998). Fecal coliforms 1. Water samples are filtered as described above. 2. The filters are placed on M-FC m e d i u m and sealed in plastic bags within 30 rain after filtration. The plates are incubated for 24 h in a water bath at 44.5°C. 4. Bacterial colonies with various shades of blue are counted as fecal coliform bacteria (APHA, 1998). LZt .
ENTEROCOCCl As stated earlier, the USEPA currently r e c o m m e n d s the use of enterococci as an indicator in marine waters. A good reference for detection and verification of enterococci is Method 1600: Membral~e Filter J~'st Method for Etlterococci ill Water, which is currently published by the USEPA (USEPA, 1997). While the USEPA r e c o m m e n d s the use of enterococci rather than the coliforms for water quality monitoring m marine waters, several points should be addressed. Like the coliforms, enterococci can be recovered in high concentrations in close association with animals, and research in ~fawaii has shown that they can survive and replicate in soils (Hardina and Fujioka, 1991). Current isolation techniques m a y be underestimating the actual numbers of enterococci in samples (Rhodes and Kator, 1997). Researchers have questioned the quality of work that led to the USEPA enterococci standards, noting that the methods e m p l o y e d by the agency to reach its conclusions m a y have been inappropriate. A short s u m m a r y of this argument, in addition to a review on the then current state of marine water quality indicators (and arguably still current) was published by Godfree et al. (1990).
"r
0L _
Equipment and reagents M e m b r a n e filtration
Same as listed for the coliforms. Enterococci media 1. m E AGAR (Difco I,aboratories, Detroit, MI) - - purchased through Fisher Scientific (Norcross, GA), product # 0333-17-9. 2. Nalidixic acid - - Fisher Scientific (Norcross, GA), product # BP908-25. 3. 2,3,5-triphenyl-tetrazolium chloride (TTC) --- Sigma (St. l,ouis, MO), product # T-8877. 4. lndoxvl [~-D-glucoside --- Sigma (St Louis, MO), product # 1-3750. 547
Enterococcus faecalis - - American Type Culture Collection (Rockville, MD), product # 19433. 6. m EI media recipe (1 1) • mEAGAR 71.2g • Distilled or deionized H20 985.0 ml • Boil to dissolve solids, autoclave, cool to 45.0°C then add • Nalidixic acid 0.24 g • 1% solution of TTC 15.0 ml (0.2 1,1mfilter sterilized) • Indoxyl [3-D-glucoside 0.81 g • Mix and then pour plates. .
Assay 1. Water samples are filtered as described using m e m b r a n e filtration. 2. The filters are placed on MEI media and incubated for 24 h at 41.0°C (Rhodes and Kator (1997) noted higher recovery w h e n incubating plates for 48 h). 3. After incubation pink or reddish colonies, which develop a blue halo are counted as enterococci (Rhodes and Kator, 1998).
CLOSTRIDIUM PERFRINGENS Clostridium pelqfringens is currently used by the state of Hawaii as an indicator to access water quality. This came about by a series of publications that demonstrated that the coliforms and enterococci were not appropriate indicators in that tropical setting (Fujioka and Shizumura, 1985; Fujioka et al., 1988; Hardina and Fujioka, 1991). While this indicator can also be recovered from animals we have typically only noted violations of the suggested 5 CFU per 100 ml in heavily polluted waters (those sites with the overall highest numbers of standard and alternative indicators). Although we typically find enterococci more widely distributed in marine waters where h u m a n viral pathogens are detected by RT-PCR, potential issues such as virus viability and detection of enterococci versus Clostridium perfrin~ens have not been resolved.
Equipment and reagents M e m b r a n e filtration
Same as listed for the coliforms. Clostridium perfringens media 1. m CP AGAR (Acumedia Manufacturers, inc., Baltimore, MD) purchased through IDEXX Laboratories, Inc. (Westbrook, ME), product # 7477. -
548
-
2. 3. 4. 5. 6. 7. 8.
9. 10.
D-cylcoserine - - Sigma (St Louis, MO), product # C-6880. Indoxy113-D-glucoside - - Sigma (St Louis, MO), product # 1-3750. P o l y m y x i n B sulfate - - Sigma (St Louis, MO), product # P-1004. Phenolphthalein diphosphate - - Sigma (St Louis, MO), product # P9875. Ferric chloride - - Sigma (St Louis, MO), product # F-2877. Clostridium perfringens - - American Type Culture Collection (Rockville, MD), product # 13124. BBL GasPak Pouch - - Becton Dickinson (Cockeysville, MD) purchased through Fisher Scientific (Norcross, GA), product # 11-81610. A m m o n i u m hydroxide - - Fisher Scientific (Norcross, GA), product # A669-500. m CP media recipe per Acumedia Manufacturers, Inc., Baltimore, MD (1 1) • mCPAGAR 71.1 g • Distilled or deionized H~O 900.0 ml • Boil to dissolve solids, autoclave, cool to 50.0°C then add • D-cycloserine 0.4 g • P o l y m y x i n B sulfate 0.025 g • 4.5% solution of ferric chloride 2.0 ml • 0.5~Xfilter sterilized solution of phenolphthalein diphosphate 20.0 ml • 0.075% filter sterilized solution of Indoxyl [3-d-Glucoside 80.0 ml • Mix and p o u r plates.
Assay 1. Water samples are filtered as described using m e m b r a n e filtration. 2. The filters are placed on m CP media and sealed with anaerobic gas paks (BBL GasPak, Becton Dickinson). 3. After 24 h incubation at 45.0°C, any yellow colonies are exposed to a m m o n i u m h y d r o x i d e fumes (do this in a fume hood) and the colonies that turn red or dark pink are counted as C. perflringens (Armon and Payment, 1988).
W-SPECIFIC R N A C O L I P H A G E Various groups have investigated the occurrence and significance of coliphage in waters being impacted by h u m a n wastes (Paul et al., 1997; Rose et al., 1997; Gantzer et al., 1998; Yanko et al., 1999). Investigations into the significance of F'-specific RNA coliphages in animal waste, h u m a n wastewater and waters u n d e r the influence of wastewater have been undertaken by a n u m b e r of researchers (Osawa et al., 1981; Havelaar et al., 1990; Debartolomeis and Cabelli, 1991; Hsu et al., 1995; Griffin et al., 2000). Tile protocol presented here was published by Hsu et al. in 1995 and contains a few modifications. We have successfully used this assay in a study 549
designed to differentiate animal from h u m a n sources of waste contamination in Florida (Griffin et al., 2000). Antibiotics are used in this assay to limit non-specific bacteria growth. The use of 2,3,5-triphenyl-tetrazolium chloride in the agar media results in a color change from tan to red w h e n catabolized by the host bacteria, which makes it easier to identify plaques. It should be stated that F+-specific RNA coliphages are a minority group of the total coliphage population. Although this assay was designed to limit the isolation of other coliphage groups, field use by this laboratory has demonstrated that a majority of the coliphage isolated, were not F--specific RNA coliphages. Isolated plaques should be verified using the prescribed protocol prior to genetic probing to limit unnecessary work. Also included is a sample enrichment assay, which was obtained from Dr Bill Yanko (Director of Wastewater, C o u n t y Sanitation Districts of Los Angeles County, San Jose Creek Water Quality Laboratory, Whittier, CA). Field studies conducted by this laboratory noted that in samples other than raw sewage or sites directly u n d e r the influence of a sewage source, the enrichment protocol had to be used to isolate F- RNA coliphage. 1. Tryptone - - Fisher Scientific (Norcross, GA), product # BP1421-500. 2. Dextrose - - Fisher Scientific (Norcross, GA), product # D16-1. 3. Sodium chloride - - Fisher Scientific (Norcross, GA), product # $271-3. 4. Agar - - Fisher Scientific (Norcross, GA), product # BP1423-2. 5. Calcium chloride - - Sigma (St Louis, MO), product # C-5080. 6.2,3,5-triphenyl-tetrazolium chloride - - Sigma (St Louis, MO), product # T-8877. 7. Streptomycin - - Sigma (St Louis, MO), product # S-2522. 8. Ampicillin - - Sigma (St Louis, MO), product # A-2804. 9. Ribonuclease A - - Sigma (St Louis, MO), product # R-4875. 10. Tris - - Fisher Scientific (Norcross, GA), product # BP152-1. 11. F o r m a l d e h y d e solution - - Fisher Scientific (Norcross, GA), product # F79-500. 12. Sodium phosphate - - Fisher Scientific (Norcross, GA), product # $374500. 13. Sodium phosphate monobasic - - Fisher Scientific (Norcross, GA), product # $369-500. 14. Lauryl sulfate - - Sigma (St. Louis, MO), product # L-5750. 15. Sodium chloride - - Fisher Scientific (Norcross, GA), product # $271-3. 16. Sodium citrate - - Fisher Scientific (Norcross, GA), product # $279-3. 17. Carnation nonfat dry milk - - Nestle USA Inc. (Solon, Ohio), can be purchased at most grocery stores. 18. I - b l o c k - Tropix (Bedford, MA), product # AL300. 19. Avidx-AP - - Tropix (Bedford, MA), product # APA10. 20. Diethanolamine (DEA) - - Tropix (Bedford, MA), product # AD120. 21. CSPD - - Tropix (Bedford, MA), product # MS050. 22. Magnesium chloride - - Sigma (St Louis, MO), product # M-1028. 23. E.coli C - - American Type Culture Collection (Rockville, MD), product # 13706. 24. E.coli Famp (F', AmpR, StrR) - - originally described and designated Escherichia coli HS(pFamp)R, by Debartolomeis and Cabelli (1991). 550
This strain of E. coli is currently being evaluated by A m e r i c a n Type Culture Collection and until available m a y be obtained from Dr M a r k Sobsey's Laboratory, University of N o r t h Carolina at Chapel Hill, L a b o r a t o r y p h o n e 1-919-966-7317. 25. Bacteriophage MS2 - - A m e r i c a n Type Culture Collection (Rockville, MD), p r o d u c t # 15597-B1. 26. Oligonucleotides - - synthesized b y O p e r o n Technologies, Inc. (Alameda, CA). 27. Media recipes for plaque overlays per H s u et al. (1995) l x t r y p t o n e broth (1 l) • 10.0 g tryptone, 1.0 g dextrose and 5.0 g NaC1 • 1 1 of distilled or deionized H~O • Boil to dissolve, autoclave and cool to 48.0°C then a d d • 5.0 ml of 3 m g ml ' a m p i c i l l i n - s t r e p t o m y c i n (mixture - - 3 m g ml ' each) Tryptone b o t t o m agar (1 1) • 10.0 g tryptone, 1.0 g dextrose, 5.0 g NaC1 a n d 12.0 g agar • 1 1 of distilled or deionized H~O • Boil to dissolve, autoclave and cool to 48.0°C then add • 5.0 ml of 3 m g ml ' a m p i c i l l i n - s t r e p t o m y c i n • 10.0 m l o f 1% TTC • For RNase A plates a d d 0.1 g 1 ~of RNase A • Mix and p o u r plates Tryptone top agar • 10.0 g tryptone, 1.0 g dextrose, 5.0 g NaC1 and 7.0 g agar • 1 1 of distilled or deionized H~O • Boil to dissolve then a d d • 0.5 ml of I N CaCL • Dispense 3.0 ml aliquots into 15.0 ml tubes and autoclave.
F-specific RNA coliphage equipment and reagents 1. Dot blot a p p a r a t u s - - Bio-Rad Laboratories (Hercules, CA) p r o d u c t # 170-6545. 2.0.45 micron M a g n a G r a p h nylon transfer m e m b r a n e - - Micron Separations Inc. (Westborough, MA), p r o d u c t # NJ4HY00010. 3. Fuji RX medical X-ray film - - Fuji Medical Systems U.S.A. inc. (Stamford, CT), p u r c h a s e d through Fisher Scientific (Norcross, GA), p r o d u c t # 04-441-97. 4. A u t o r a d i o g r a p h y cassette - - Fisher Scientific (Norcross, GA), p r o d u c t # FB-XC-810. 5. K o d a k GBX film d e v e l o p e r / r e p l e n i s h e r and f i x e r ~ r e p l e n i s h e r -Eastman K o d a k C o m p a n y (Rochester, NY), p r o d u c t # CAT-190-1859. 6. UV crosslinker - - Fisher Scientific (Norcross, GA), p r o d u c t # FBUVXL-1000. 7. Southern-Light C h e m i l u m i n e s c e n t Nucleic Acid Detection Protocol (biotin labeled probes) - - Tropix (Bedford, MA) - - can be d o w n l o a d e d from www.tropix.com.
$51
8.15 ml p o l y p r o p y l e n e tubes - - Corning Incorporated (Corning, NY), purchased through Fisher Scientific (Norcross, GA), product # 05-53859A. 9. 100 m m x 15 m m disposable Petri dishes - - Fisher Scientific (Norcross, GA), product # 08-757-13. 10. Hybridization bags - - Life Technologies, GIBCO BRL Products (Rockville, MD), product # 18278-010. 11. P o u c h / b a g sealer - - Kapak Corporation (St Louis Park, MN), product # 101-1. 12. Miscellaneous e q u i p m e n t - - microwave, 65.0°C incubator, 45.0°C incubator, 48.0°C waterbath, 15 ml tube racks, darkroom, Pasteur pipettes, 1.5 ml microfuge tubes, micropipetters.
Methods Enrichment assay 1. The host is g r o w n overnight in 10.0 ml of tryptone broth (1 1 of l x tryptone broth - - 10.0 g tryptone, 1.0 g dextrose, 5.0 g NaC1) containing 15.0 gg ml ~ampicillin-streptomycin at 35.0°C. 2. Three hours prior to the start of the assay, l m l of host from the overnight culture is inoculated into 49.0 ml of fresh tryptone broth. 3. 1 1 grab samples then are inoculated with 10.0 ml of host and 100.0 ml of 11x tryptone broth. 4. The samples are then incubated for 48 h at 35.0°C (no shaking). 5. After the 48 h, 1.0 ml of host is a d d e d to 3.0 ml of top agar (1 1 of l x top agar - - same as l x tryptone broth without the antibiotics and with 1.0 ml of 1.0 N CaCI~), which has been melted and cooled to 48.0°C. 6. The top agar and host are then poured over a plate of tryptone bottom agar (11 of l x bottom agar - - same as l x tryptone broth) containing 15.0 ~g m l ' ampicillin-streptomycin and 0.01% 2,3,5 triphenyl tetrazolium chloride. 10 gl of the enriched sample is then spotted onto the host plate. 7. The plates are incubated overnight at 35.0°C. 8. 1 ml dilutions to 10 7were completed on all samples (from the enriched 1 1 grab samples), which produced plaques. 9. The 1.0ml dilution sets are then utilized for overlays as described below.
F RNA coliphage assay 1. The host (E. coli Famp) is grown overnight in 10.0 ml of l x tryptone
broth at 35.0°C. . Three hours prior to the start of the assay, 1 ml of host from the
overnight culture is inoculated into 49.0 ml of fresh tryptone broth and incubated at 35.0°C. . After 3 h, 1.0 ml of host and 1.0 ml of sample (grab sample or enriched sample dilutions) is a d d e d to 3.0 ml of top agar (which is liquified in a boiling water bath or microwave and cooled to 48.0°C prior to use). 552
4. The top agar, sample and host are mixed and then poured over a plate of tryptone bottom agar (this may be repeated to screen a desired volume of sample. Typically 5.0 to 10.0 ml of sample is assayed using this technique). 5. A negative control overlay is made by using top agar, host and distilled H20. A positive control overlay is made by using top agar, host and distilled H~O, pouring the overlay and after the overlay has solidified, placing a 5.01al drop of concentrated MS2 phage in the center of the plate. 6. After the plates solidify, they are incubated for 24 h at 35.0°C. 7. After 24 h plaque forming units (pfu) are counted. Plaques appear as a cleared zone, pinprick in size on the red bacterial lawn. The cleared area on the positive control will be as large as the 5.0 lal drop of MS2 (F÷-specific RNA coliphage control). A desired number of plaques can then be picked from each plate and screened for differentiation (is the plaque really a plaque and if so is the phage a DNA somatic or F ' RNA?) prior to genetic probing. If the numbers of plaques per plate are large we typically screen 20 per plate. However, the greater the number of plaques screened, the greater the chance of detecting Fspecific coliphages of human origin, which are usually rare in the environment. 8. Plaques are picked using a Pasteur pipette and placed in 1.0 ml of 0.5 M Tris (pH -8.0). Plaques are then vortexed briefly (plaques may be stored at 4.0°C for up to 1 week until further analysis). 9. Plaques were then verified by spotting 5.0pl of the TRIS/plaque suspension onto a fresh lawn of E. coli Famp (from 3 h culture) using tryptone bottom agar media; 5.0 lal onto a lawn of E. coil Famp using tryptone bottom agar containing RNase (to control for DNA coliphage); 5.0 I~1 onto a lawn of E. coli C using tryptone bottom agar (no antibiotics) with and without RNase A. To limit the number of agar plates used, the plates may be subdivided using a permanent marker (typically 4 to 6 sections per plate). 10. Plates are then incubated for 24 h at 35.0°C (do not invert plates). 11. Plaque forming units are identified as follows: • Plaque forms on the E. coil Famp/bottom agar plate w / o RNase A and not on the remaining plates = F RNA coliphage. • Plaque forms on the E. coli F a m p / b o t t o m agar plates with and w / o RNase A and not on the remaining plates = rare but observed, which would indicate an F ~DNA coliphage capable of lysis. • Plaques form on all plates = somatic DNA phage. • Plaque forms on the E. coli Famp/bottom agar plate w / o RNase A and on the E. coli C/bottom agar plate w / o RNase A and not on any of the remaining plates = rare but observed, which would indicate a somatic RNA phage. • No plaques on any of the plates = not a phage. 12. Those plaque forming units identified as F" RNA coliphage can then be screened with the genetic probes to determine their source using the following protocol.
553
¢4-/ m
"1" uP.
O u .B
Genotyping F* RNA coliphages using oligonucleotide probes (modified assay from Hsu et al., 1995) 1. Isolates that are identified as F ~RNA coliphage are picked (from the verification/differentiation plate not the original isolate) using a Pasteur pipette. A total of ten plugs per viral clearing are picked and suspended in a buffer containing 537.0 pl of 20x SSC (3.0 MNaC1, 0.3 M Na-citrate, pH 7.0), 496.0pl of 37q~, formaldehyde and 400.01ll of distilled water (which is irradiated with UV light using the optimal cross-link setting on the Fisher Scientific UV cross-linker). N o t e - - individual plaque lifts do not form sufficient hybridization signal. 2. These suspensions are then incubated at 65.ff'C for 30.0 rain. 3. Aliquots of 335.0 1-11(×4, one for each virus probe group) are then applied to 0.45 ~ml positively charged nylon filter paper using a dot blot apparatus. 4. Fix the viral RNA to the filters using UV light (optimal cross-link setting on the Fisher UV cross-linker). 5. Filters are then cut into four sections (one section per probe(s)). The probes (Table 27.2, sequences per Hsu e t a l . , 1995) for detection of group Ii and group III F÷ RNA coliphages (found predominately in human feces) and for the detection of group I and group IV F RNA coliphages (found predominately in animal feces) are employed for hybridization and detection. 6. Hybridization and stringency wash protocols and reagents are per TROPIX Inc. Southern-Light Chemiluminescent Nucleic Acid Detection Protocol (all reagents needed have been listed). Use non-fat dried milk as a blocking reagent (1.0%) in both the prehybridization and hybridization solutions. 7. Blots are imaged with X-ray film (typical exposure time is 1 h).
Table 27.2 F RNA coliphage probe sequences Virus
Group Probe sequence
MS2 GA
I IIa IIb III IV
QB SP/FI
Target region
5'-CTAAGGTATGGACCATCGAGAAAGGA-3' 5'-CATGTTATCCCCCAAGTGCTGGCTAT-3' 5'-GTTTTCCTTATGTTTTGCTTTCAGACCCA-3' 5'-ATACTCAGTGAA(A/G)TACTGCTGTGT-3' 5'-GGCATAGATTCTCCTCTGTAGTGCG-3'
Maturation protein Maturation protein 5' nontranslated region 5' nontranslated region
CONCLUSIONS Currently there is no ideal indicator for all marine environments and active screening of marine samples for multiple indicators may be required to determine a region's best available indicator. Molecular techniques such as RT-PCR, PCR and immuno-fluorescent assays may be employed for pathogen screening in addition to screening for indicators. In Chapter 28 of this volume, several molecular methods for pathogen 554
detection are presented, w h i c h h a v e p r o v e d v a l u a b l e in d i s c e r n i n g the best available indicator in Florida's m a r i n e waters. Of particular use has b e e n screening s a m p l e s for the p r e s e n c e of h u m a n e n t e r o v i r u s e s a n d Hepatitis A viruses. Since h u m a n s are the o n l y k n o w n host of Hepatitis A viruses a n d e n t e r o v i r u s e s such as the polioviruses, screening of s a m p l e s for their p r e s e n c e has a l l o w e d us to d e t e r m i n e if h u m a n w a s t e s or a n i m a l w a s t e s are i m p a c t i n g b o d i e s of water. O t h e r p a t h o g e n s such as the N o r w a l k viruses ( S c h w a b et al., 1997) a n d Small R o u n d S t r u c t u r e d viruses ( A n d o et al., 1995; F a n k h a u s e r et al., 1999) in a d d i t i o n to t e c h n i q u e s such as b i o m a r k e r profiling (Cajaraville et al., 2000) a n d antibiotic resistance profiling (Parveen et al., 1997), m a y p r o v e to be v a l u a b l e tools in identif y i n g the ideal indicator(s) or indicator system.
References Ando, T., Monroe, S. S., Gentsch, ]. R., Jin, Q., Lewis, D. C. and Glass, R. I. (1995). Detection and differentiation of antigenically distinct small round-structured viruses (Norwalk-like viruses) by reverse transcription-PCR and Southern hybridization. J. CliJz. Microbiol. 33, 64-71. APHA (American Public Health Association) (1998). Standard Methods for the Examillatioll of Water alu/ Wastewater, 20th edn. American Public Health Association, Washington, DC. Armon, R. and Payment, P. (1988). A modified m-CP medium for enumeration Clostridium pclfril~gens from water samples. Cat1. J. Microbiol. 34, 78-79. Byrd, J. J. and Colwell, R. R. (1993). Long-term survival and plasmid maintenance of Escherichia coil in marine microcosms. FEMS Microbial Ecol. 12, 9-14. Cabelli, V. J. (1983). Health effects for marine recreation waters. Health Effects Research Laboratory. Research Triangle Park, NC. USEPA 600/1-80-031. Cajaraville, M. P., Bebianno, M. J., Blasco, J., Porte, C., Sarasquete C. and Viarengo, A. (2000). The use of biomarkers to assess the impact of pollution in coastal environments of the Iberian Peninsula: a practical approach. Sci. Total Environ. 247, 295-311. Debartolomeis, I. and Cabelli, V. J. (1991). Evaluation of an Escherichia coli host strain for enumeration of male-specific bacteriophages. Appl. EHviron. Microbiol. 57, 1301-1307. Dufour, A. P., Ericksen, T. H., Ballentine, R. K., Cabelli, V. J., Goldberg, M. and Fox, W. E. (1986). Bacteriological ambient water quality criteria for marine and fresh recreational waters. United States Environmental Protection Agency. Ambient Water Quality Criteria for Bacteria. EPA44075-84-002. Escherich, T. (1885). Die Darmbacterien des Neugeberenen und Sauglings Fortschritte der Medicin 16, 515 524. Fankhauser, R. L., Noel, J. S., Monroe, S. S., Ando, T. and Glass, R. I. (I999). Molecular epidemiology of 'Norwalk-like viruses' in outbreaks of gastroenteritis in the United States. J. Infectious Diseases 178, 1571-1578. Fujioka, R. S. and Shizumura, L. K. (1985). Clostridium perfrillgel~s a reliable indicator of stream water quality. J. Water Po/lutioH CoJ#ro[ Fed. 57, 986-992. Fujioka, R. S., Tenno, K. and Kansako, S. (1988). Naturally occurring fecal coliforms and fecal streptococci in Hawaii's freshwater streams. Toxicity Assessment 3, 613-630. Gantzer, C., Maul, A., Audic, J. M. and Schwartzbrod, L. (1998). Detection of infectious enteroviruses, enterovirus genomes, somatic coliphages, and Bacteroides hagilis phages in treated wastewater. Appl. El~viroH. Microbiol. 64, 4307-4312.
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Garcia-Lara, J., Menon, P., Servais, P. and Billen, G. (1991). Mortality of fecal bacteria in seawater. Appl. Environ. Microbiol. 7, 885-888. Gerba, C. P. and Mcleod, J. S. (1976). Effect of sediments on the survival of Escherichia coli in marine waters. Appl. Environ. Microbiol. 32, 114-120. Gleeson, C. and Gray, N. (1997). The Coliform Index and Waterborne Disease. E & FN Spon, London. Godfree, A., Jones, E and Kay, D. (1990). Recreational water quality. The management of environmental health risks associated with sewage discharges. Mar. Pollution Bull. 21,414-422. Griffin, D. W. (1999). Microbiological Studies of Florida Water Quality. Dissertation. University of South Florida, St Petersburg, Florida. Griffin, D. W., Gibson C. J. III, Lipp, E. K., Riley, K., Paul, J. H. and Rose, J. B. (1999). Detection of viral pathogens by reverse transcriptase PCR and of microbial indicators by standard methods in the canals of the Florida Keys. Appl. Environ. Microbiol. 65, 4118-4125. Griffin, D. W., Stokes, R., Rose, J. B. and Paul, J. H. (2000). Bacterial indicator occurrence and the use of an F specific RNA coliphage assay to identify fecal sources in Homosassa Springs, Florida. Microbial Ecol. 39, 56-64. Hagler, A. N., Rosa, C. A., Morris, P. B., Mendonca-Hagler, L. C., Franco, G. M. O., Araujo, E V. and Soares, C. A. G. (1993). Yeasts and coliform bacteria of water accumulated in bromeliads of mangrove and sand dune ecosystems of southeast Brazil. Can. J. Microbiol. 39: 973-976. Hardina, C. M. and Fujioka, R. S. (1991). Soil: the environmental source of Escherichia coli and Enterococci in Hawaii's streams. Envin~n. Toxicol. Water Quality 6, 185-195. Havelaar, A. H., Pot-Hogeboom, W. M., Furuse, K., Pot, R. and Hormann, M. P. (1990). F-specific RNA bacteriophages and sensitive host strains in faeces and wastewater of human and animal origin. ]. Appl. Bacteriol. 69, 3960-3966. Hazen, T. C. and Toranzos, G. A. (1990). Tropical source water. In Drinking Water Microbiology (G.A. McFeters, Ed.), pp. 32-53. Springer-Verlag, New York. Huanca, W., Santander, E., Padilla, L. and Mondaca, M. A. (1996). Survival of gram negative bacteria in seawater. Gayana Oceanologia 4, 153-157. Hsu, F-C., Shieh, Y-S. C., van Duin, J., Beekwilder, M. J. and Sobsey, M. D. (1995). Genotyping male-specific RNA coliphages by hybridization with oligonucleotide probes. Appl. Environ. Microbiol. 61, 3960-3966. Lapointe, B. E., O'Connell, J. D. and Garrett, G. S. (1990). Nutrient couplings between on-site waste disposal systems, groundwater, and nearshore surface waters of the Florida Keys. Biogeochent. 10, 289-307. Lipp, E., Farrah, S. and Rose, J. (1997). A Study on the Presence of Human Viruses in Surface Waters of Sarasota County. Sarasota, Florida Department of Health Water Quality Report. Lipp, E., Kurz, R., Vincent, R., Rodriguez, C. and Rose, J. (1999). Assessment of the Microbiological Water Quality of Charlotte Harbor Florida. Souffi-West Florida Water Management District Technical Report. Lisle, J. (1993). An Operator's Guide to Bacteriological Testing. American Water Works Association, Denver, CO. NRDC (National Research Data Center) (1998). Testing the Waters. Osawa, S., Furuse, D. and Watanabe, I. (1981). Distribution of ribonucleic acid coliphages in animals. Appl. Environ. Mictvbiol. 41, 164-168. Parveen, S., Murphree, R. L., Edmiston, L., Kasper, C. W,, Portier, K. M. and Tamplin, M. L. (1997). Association of multiple-antibiotic-resistance profiles with point and nonpoint sources of Escherichia coil in Apalachicola Bay. Appl. Envirol~. Microbiol. 63, 2607-2612.
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Paul, J. H., Rose, J. B., Jiang, S., Kellogg, C. and Shinn, E. (1995). Occurrence of fecal indicator bacteria in surface waters and the subsurface aquifer in Key Largo, Florida. Appl. Environ. Microbiol. 61, 2235-2241. Paul, J. H., Rose, J. B., Jiang, S. C., Zhou, X., Cochran, P., Kellogg, C., Kang, J. B., Griffin, D., Farrah, S. and Lukasik, J. (1997). Evidence for groundwater and surface marine water contamination by waste disposal wells in the Florida Keys. Water Res. 31, 1448-1454. Perez-Rosas, N., and T. C. Hazen, (1988). In situ survival of Vibrio cholerae and, Escherichia coli in tropical coral reefs. Appl. Dzviron. Microbiol. 54, 1-9. Rhodes, M. W. and Kator, J. (1997). Enumeration of EHterococcus sp. using a modified mE method. J. Appl. Microbiol. 83, 120-126. Rose, J. B., Zhou, X., Griffin, D. W. and Paul, J. H. (1997). Comparison of PCR and plaque assay for detection and enumeration of coliphage in polluted marine waters. Appl. Eilviron. Microbiol. 63, 4564-4566. Santiago-Mercado, J. and Hazen, T. C. (1987). Comparison of four membrane filter methods for fecal coliform enumeration in tropical waters. Appl. EJTviropl. Microbiol. 53, 2922-2928. Schwab, K. J., Estes, M. K., Neill, E H. and Atmar, R. L. (1997). Use of heat release and an internal RNA standard control in reverse transcription-PCR detection of Norwalk virus from stool samples. J. Cli~1. Microbiol. 35, 511-514. USEPA (United States Environmental Protection Agency) (1997). Method 1600: Membrane Filter Test Method (or Enterococci in Water. Washington, DC, EPA-821R-97-004. USEPA (United States Environmental Protection Agency) (1998). Bacterial Water Quality Standard for RecreatioHal Waters (Freshwater and Mari~Te Waters). Washington, DC, EPA-823-R-98-003. Yanko, W. A., Jackson, J. L., Williams, E P., Walker, A. S. and Castillo, M. S. (1999). An unexpected temporal pattern of coliphage isolation in groundwaters sampled from wells at varied distances from reclaimed water recharge sites. Water Res. 33, 53-64.
List of suppliers AAPER Alcohol and Chemical Co. P.O. Box 339 Shelbyville KY 40066-0339, LISA Teh 1-800-456-1017 Website: http://www.aaper.com American Type Culture Collection 12301 Parklawn Drive Rockville M D 20852-1776, USA Teh 1-800-638-6597 Website: http://ww~ .atcc.org
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Bio-Rad Laboratories 2000 Alfred Nobel Drive Herctdes, CA 94547, USA Teh 1-800-424-6723 Website: http://www.bio-rad.com Eastman Kodak Company 343 State St~vet Rochester N Y 14650, LISA Teh 1-800-242-2424 Website: http://www.kodak.com
Fisher Scientific P.O. Box 4829 Norcross, GA 30091, USA Tel: 1-800-766-7000 Website: http://www.fishersci.com
Operon Technologies, Inc. 1000 Atlantic Avenue Alameda, CA 94501, USA Teh 1-800-688-2248 Website: http://www.operon.com
IDEXX Laboratories, Inc. One IDEXX Drive Westbrook ME 04092, USA Tel: 1-800-321-0207 Website: http://www.idexx.com/fed/ products/acumedia.asp
Pall Gelman Sciences 600 South Wagner Road Ann Arbor MI 48103-9019, USA Tel: 1-800-521-1520 Website: http://www.gehnan.com
Kapak Corporation 5303 Parkdale Drive St Louis Park M N 55416-1681, USA Tel: 1-800-527-2557 Website: http://www.kapak.com Life Technologies, GIBCO BRL Products 9800 Medical Center Drive P.O. Box 6482 Rockville, MD 20849, USA Tel: 1-800-828-6686 Website: http://www.lifetech.com Micron Separations Inc. 135 Flanders Road Westborough, MA 01581, USA Tel: 1-800-444-8212 Website: http://www.msifilters.com
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SIGMA P.O. Box 14508 St Louis MO 63178, USA Teh 1-800-325-3010 Website: http://www.sigmaaldrich.corn Tropix 47 Wiggins Avenue Bedford MA 01730, USA Tel: 1-800-542-2369 Website: http://www.tropix.com
28 H u m a n Enteric Viruses and Parasites in the Marine Environment Erin K L i p p ' , J e r z y L u k a s i k 2 and Joan B Rose 3
'Center of Marine Biotechnology, University of Maryland Biotechnology Institute, Baltimore, Maryland, USA; 2Biological Consulting Services of North Florida, Inc., Gainesville, Florida, USA; 3College of Marine Science, University of South Florida, St. Petersburg, Florida, USA
CONTENTS Introduction Enteric viruses Enteric protozoan parasites Methodological issues
eeeeee
INTRODUCTION Oceanic and coastal waters are k n o w n to harbor and transport microorganisms that cause disease in h u m a n s and other animals. Additionally, as modulators of climate the oceans m a y indirectly influence disease patterns and the distribution of m a n y pathogens. While certain pathogenic or toxigenic microorganisms, including toxic p h y t o p l a n k t o n and Vibrio spp., occur naturally in marine and estuarine waters, anthropogenic microbes including enteric bacteria, protozoa and viruses m a y be introduced to coastal waters in sewage pollution. Despite the relatively unfavorable environment, these introduced organisms m a y survive for prolonged periods in the marine environment, often associated with sediments and other protective particles (LaBelle and Gerba, 1982). Risk to swimmers using polluted beaches has been the major determinant in establishing ambient water quality standards and discharge limits to recreational sites. In addition, shellfish-associated disease remains a significant public health concern (Lipp and Rose, 1997). Public health issues for marine waters in the tropics and subtropics differ from those of cooler waters. Thus there is geographical distinction for these environments t h r o u g h o u t the world and the current system for assessing marine water quality and public health risks is inadequate. Prevention of disease d e p e n d s on appropriate monitoring strategies. Indeed, more emphasis is n o w being placed on direct monitoring for pathogens of concern.
METHODSIN MICROBIOLOGY,VOLUME30 ISBN 0-I 2 521530-4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any form reserved
ENTERICVIRUSES Background Water is a common vehicle for the transmission of many enteric viruses (Goyal 1984; Gerba et al., 1985). More than 100 viruses have been identified in human feces including Hepatitis A, enteroviruses and Norwalk viruses. Illnesses caused by enteric viruses include fever, rash, meningitis, paralysis, myocarditis, hepatitis, eye infections, gastroenteritis, flu-like symptoms and chronic fatigue syndrome (Melnick and Gerta, 1980; 1984; Muir et al., 1998). In surface water the primary source of enteric viruses is feces (human, bovine or swine) which may originate from storm water and treated or untreated wastewater. A major route for enteric virus entry into estuaries is through tributary streams, rivers and canals that have been impacted by wastewater. Septic tanks and sewage treatment facilities are of particular concern in coastal urban areas. In the Florida Keys, bacteriophages seeded into septic tanks were able to move rapidly through the subsurface to nearby canals and offshore waters (Paul et al., 1997). Moreover, widespread virus contamination has been detected in the Florida Keys canals by RT-PCR (Griffin et al., 1999) and in Charlotte Harbor in shellfish areas open to harvesting during the rainy season (Lipp et al., in press (a)). Viruses present a special problem in terms of their potential for contamination. Their small size and resistant protein coat allow for easier transport and prolonged survival in the environment. Enteric viruses survive longer in fresh and marine water than coliform bacteria, which are used to monitor water quality (Bitton et al., 1983; Craun, 1984; Payment et al., 1985; Havelaar et al., 1993). Additionally, viruses in coastal waters have been detected far from the original source of pollution and in the absence of bacterial indicators (Metcalf and Stiles, 1967; Lipp et al., in press (a)).
Collection of enteric h u m a n viruses from m a r i n e waters Principle and application In marine and other surface waters, numbers of enteric viruses are often too low to be detected in unconcentrated samples. Therefore, large volumes of water must be concentrated by adsorption/filtration and elution (desorption) before analysis. Due in part to the small size of an individual viral particle, charge interactions are optimized to retain enteroviruses on electrostatic filters, rather than mechanical capture. Currently, both electronegative and electropositive filters are available; they each have certain advantages and disadvantages. Because most enteric viruses are negatively charged at ambient conditions they can adsorb directly to an electropositive filter (i.e. 1MDS Virosorb; CUNO, Meriden, CN). However, these filters clog relatively easily, they are not effective when the pH is above 8.0, and they have a poor recovery rate for
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viruses in marine waters. Electronegative filters have a greater capacity for virus adsorption in marine waters and waters of high turbidity, organic matter and pH. However, they require substantial pre-conditioning of the water to facilitate virus adsorption. The most commonly used type of electronegative filter is the Filterite filter (Filterite Corp., Timonium, MD). Because of its superior efficiency in marine waters, the virus concentration procedure described in the following section is based on the electronegative filter (Fig. 28.1).
Adsorption/filtration
Materials and reagents • 100-200 I carboy or plastic trash can (30-50 gallons) • Small gas powered water pump (i.e. Homelite or Tanaka) with screened, I inch diameter intake hose (generally provided with pump) • 30-50 cm of pressurized hose (~ inch diameter) with metal hose clamps to hold male and/or female hose fittings (preferably the type that can snap together) • Cartridge housings for 10 inch filters fitted with appropriate hose fittings (see above) • Volume meter with 0.10 gallon resolution fitted with appropriate hose fittings • Portable temperature-compensated pH meter (small battery-operated type) • Hydrochloric or acetic acid (3-5 M) • Sodium hydroxide ( I - 4 M) • Aluminum chloride (3 M) • 25 ml sterile pipettes and bulb Procedure
1.
2.
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Collect the sample into a large pre-bleached container (minimum 100 1 capacity) using a gasoline-driven or electric pump. Generally between 100 and 200 1 can be easily collected in 30 or 50 gallon trashcans. Adjust the pH to 3.5 using a 3-5 M solution of hydrochloric or acetic acid. The isoelectric point of most enteric viruses occurs at a pH of N4.0. The pH must be below this point to ensure that the virus particles are positively charged and therefore capable of adsorbing to the electronegative filter. The sample should be mixed continuously (this can be done be recirculating the water through the pump system). Add 0.5 ml of 3 M aluminum chloride (A1CI~) per liter of collected water, the final concentration of aluminum chloride should be 1.5 raM. This results in the flocculation of organic matter in the surface water, which helps to minimize viruses binding to other organic material.
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4.
5. 6.
After appropriate conditioning, the p u m p should be turned off. The outflow hose should then be connected to a cartridge housing containing a filter. (The housing must be able to hold a 10 inch cartridge filter.) A v o l u m e meter should be connected after the housing to allow the determination of both flow rates and total v o l u m e filtered. Once all connections are made, the entire v o l u m e should be p u m p e d out and filtered at a flow rate _<30 1 rain 1.
Elution
Materials and reagents • • • •
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N2 tank and pressure vessel OR source of compressed air Length of pressurized hose Sterile I I polypropylene bottles Sterile 0.5-1.5% beef extract with 0.5 M glycine (BE/G) at pH 9.0-9.5.The preferred beef extract is Beef Extract V from BB/ (Becton Dickinson), which can be used at the higher concentrations. However, recently the company stopped production. If BE V is not available, desiccated beef extract (Difco) should be used at 0.5% concentration.This type of BE does not flocculate as well as the BEV and therefore further manipulation with FeCI 2 during the concentration step is required (Payment et al., 1984) NaOH (I M) Hydrochloric or acetic acid (I M) Temperature compensated pH meter
Procedure
1.
2. 3. 4.
5.
Viruses are desorbed or eluted from the filters by reversing the charge on the protein coat. This can be accomplished by adding a high p H beef extract solution buffered with 0.5 M glycine (BE-G, p H 9). Sterile BE-G (1 1) can be either poured over the filter in the housing and forced out using compressed air, or a pressure vessel can be used to push the BE-G through the filter and housing. In either case, the eluent should be carefully collected in a sterile beaker or p o l y p r o p y l e n e container. Repeat step 1. Neutralize the eluate with 1 M HC1 or acetic acid and record final volume. Eluate can be stored 4°C for 24 h until further concentration and processing. For long-term storage, the eluate can be frozen at -70°C. To avoid contamination between samples, all equipment should be disinfected with a 10% bleach solution).
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Concentration of eluate
The p r o c e d u r e for concentrating the filter eluate is a d a p t e d from the USEPA's Information Collection Rule (ICR) protocol (USEPA, 1995).
Materials and reagents
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• • • • • • • •
Refrigerated centrifuge with 2500-I 0 000g capability (must be able to hold 100-1000 ml screw cap bottles) Syringe-driven 0.22pm sterilizing filter, i.e. Costar (pre-exposed to 10-20 ml of beef extract at pH -7.0) OR 0.22 lum low protein binding durapore (PVDF) filter without beef extract treatment (i.e. Millepore brand Millex HV) Sterile 10-60 cc syringes Water bath set at 37°C Stir bars (sterilized) and magnetic stirrers Screw cap sterile centrifuge bottles (I 00-500 ml) 0.15 M solution of Na2HPO4"7H~O (sterile) Hydrochloric acid (I M) NaOH (I M) Ferric chloride
Procedure
1. M a k e u p a 0.15 M solution of Na2HPO~-7H~O, adjust to p H 9.0-9.5 and filter sterilize through a 0.22 p m pre-beef extract treated filter. Alternatively a 0 . 2 2 p m low protein binding d u r a p o r e filter (PVDF) can be used w i t h o u t beef extract treatment (i.e. Millex ~ GV). 2. If eluate was stored at -70 °, defrost in a 37 ° w a t e r bath. 3. If Difco desiccated beef extract was used, sterile ferric chloride should be a d d e d to a final concentration of 0.25 mM to p r o m o t e flocculation ( P a y m e n t et al., 1984). 4. Slowly a d d 1 N HC1 to the t h a w e d or fresh s a m p l e until the solution reaches a p H of 3.5 + 0.1. 5. Stir s a m p l e slowly on a magnetic stirrer for 30 rain at r o o m temperature. 6. Pour the contents into 100-500ml capacity centrifuge bottles (several m a y be needed). Centrifuge at 4°C for 15 min at 2500g. 7. Discard supernatant. 8. Place a stir bar in centrifuge bottles containing the precipitate and stir with 30-50 ml of 0.15 M Na2PO4.7H:O for 20 minutes or until the precipitate has dissolved. 9. Readjust the p H to 9.0-9.5 if necessary, r e m o v e the stir bar and centrifuge again at 4000-10 000~ for 10 min at 4°C. 10. Save the s u p e r n a t a n t a n d adjust p H to 7.0-7.5 with 1 M HC1. To reduce bacterial contamination, force the s u p e r n a t a n t through a syringe-driven low protein binding d u r a p o r e filter (PVDF), 0.22 p m (i.e. Millex'" - GV).
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11. The final volume of supernatant should be recorded. The amount of final concentrate assayed can then be back calculated to the volume of original source water collected.
Troubleshooting The adsorption-elution-concentration procedures are relatively straightforward and rarely require additional manipulation. However, occasional problems may arise. Throughout all procedures, care should be taken to keep pH at the desired range for each step. Particularly, pH below 3.0 may inactivate viruses. Because of this, the elution step should be carried as soon as possible after collection, although a holding time of up to 6 h is acceptable if the filter is kept at 4°C. In the collection/adsorption phase, turbid samples may clog the charged filter before the desired volume has been collected. In these cases a 1 Ixm nominal porosity cartridge pre-filter can be placed in line before the charged filter to remove larger particles. Ideally this filter should also be assayed for any particle-bound viruses. In the later elution and concentration phases it is important to minimize the amount of foam produced in the beef extract, which will inactivate viruses. This is most often a problem at the elution step. Simply reduce the pressure to prevent excessive foaming. During elution and concentration it is also imperative to decontaminate the pH meter electrode between samples. In the field this is easily accomplished by immersion in 100% bleach solution.
Detection of human viruses by cell culture Principle and applications While more than 100 types of human pathogenic viruses may be present in fecally contaminated water, only a few can be detected by currently available methods. Of these methods, those that rely on concentration, elution, and then detection on cell culture have gained acceptance. These methods usually detect enteroviruses and reoviruses; the detection of adenoviruses, hepatitis A virus, astroviruses and rotaviruses by cell culture is possible but methods are tedious and less often used. Therefore, water quality evaluation has come to rely on detection of culturable enteroviruses and reoviruses. The standard method for the detection of enteroviruses as mandated by the Environmental Protection Agency (EPA) involves the use of the Total Culturable Virus Assay. The principle relies on the availability of cell strains that are susceptible to, and support the replication of human enteric viruses. In addition, the infected cells must also develop cytolytic morphological changes that can be visually determined. This observation is commonly termed cytopathogenic effects (CPE). Approved by the EPA, the Buffalo Green Monkey (BGM) cell line is the most commonly used for the detection of enteric viruses. BGM cells
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compose a continuous cell line derived from African Green Monkey kidney cells. They are highly susceptible to many enteric viruses (Dahling et al., 1984; Dahling and Wright, 1986) and have been used to recover various enteroviruses from environmental samples (Morris and Waite, 1980; Schmidt et al., 1978). The characteristics of this line have been previously described by Barron et al. (1970). Many researchers have also used other cell strains for the detection of enteroviruses from water. The strains used include RD, CaCo-2, and MA104. Our lab has found these cell strains to have increased detection sensitivity to various viral pathogens, especially rotaviruses, reoviruses and astroviruses. Regardless of the cell strain used for viral detection, the passage number must be relatively low (usually below 200). It has been demonstrated that the higher the passage number the less sensitive the cells are to viral infection. It should be noted, however, that currently only some viruses can be detected by cell culture. Other viruses such as SRSV (Small Round Structured Viruses), Rotavirus, Astrovirus, etc. do not exhibit CPE in cell culture. Consequently, other methods should be used for their detection (such as integrated cell culture-PCR/RT-PCR, see below).
Materials and reagents • • •
•
•
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Incubator capable of maintaining the temperature of cell cultures at 36.5 + I°C. Inverted microscope capable of 40-100× magnifications. Class II biological safety hood or a clean area dedicated to tissue cell culture work.The hood or area must be separated from the one used to pass and generate cell stocks. Sterilizing filter - - 0.22 IJm (Costar Product No. 140666). Always pass about 10-20 ml of 1.5% beef extract, pH 7.0-7.5, through the filter just prior to use to minimize virus adsorption to the filter. Cell culture flasks (Coming Costar Inc., 25 cm ~ flasks part number 430372) containing a confluent monolayer of BGM cells (American Type Culture Collection) or any other susceptible animal-derived cell line. Cell cultures used for viral assays are most sensitive to viral infection between 48 and 96 h following their most recent passage; cells older than 7 days should not be used. Sterile, disposable I ml, 5 ml and 25 ml pipettes. Sterile 0.15 t4 Na~HPO4- 7H20 adjusted to pH 7.0-7.5. Eagle Minimum Essential Medium (MEM) w/ Earle's salts, L-glutamine (Mediatech product number 10-010-CM) adjusted to 2% Fetal Bovine Serum (Mediatech product number 35-010-CV), 100u ml ' penicillin G, 100 lug ml' Streptomycin, and 0.25 IJg ml' Amphotericin B (Mediatech product number 30-004-CI).
Procedure
All the following steps must be performed in a biological safety hood. It is recommended that the hood be clean and disinfected first by either
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UV radiation or wiping with 70% ethyl alcohol. If a hood is not available, the work should be performed in a clean disinfected area (with ethyl alcohol) that is designated for tissue culture inoculation. Aseptic technique must be used throughout, and caution must be exerted w h e n handling the environmental viral concentrates and used cell culture media. Touching either the cannula or the pipetting device to the inside rim of the cell culture test vessels should be avoided to avert the possibility of transporting contaminants to the remaining culture vessels. 1. The cell on flasks should first be examined microscopically to ensure the presence of confluent monolayers. 2. Media of the cell culture flasks is decanted into a sterile beaker gently to avoid splashing. 3. A subsample of the collected concentrate is rapidly thawed. Usually only 1/20th of the assay sample v o l u m e is inoculated initially to ensure the absence of contamination and toxicity problems. Initial inoculation is usually done onto ten 25 cm-' cell culture flasks. The virus sample is inoculated onto a media-free monolayer of cells. The m a x i m u m v o l u m e of inoculated sample is d e p e n d e n t on available cell surface area. Inoculum v o l u m e should not exceed 0.04 ml cm -"of the flask's monolayer surface area. If viral concentrate is expected to contain high concentration of infective viruses, then five-fold dilutions of the sample should be m a d e in sterile 0.15 M NaeHPO~. l ml of 0.15 M Na_~HPOJH~_O is inoculated in a separate flask containing a monolayer of cells. This will serve as the negative control. 4. The flasks are sealed, laid horizontally and intermittently rotated slowly so that the sample is allowed to incubate onto the cell for 120 min. This will permit virus adsorption to the cells. 5. After incubation, MEM w / 2 % FBS nutrient media p r e - w a r m e d to 36.5 _+ I°C is a d d e d to each flask. Sufficient solution is a d d e d to ensure the covering of the whole monolayer surface area. This is usually equivalent to one-third of the indicated growth surface area of the flask. The solution should be a d d e d to the side of the cell culture vessel opposite the cell monolayer in order to avoid disturbing the monolayer. 6. The flask is incubated at 36.5 _+ 1°C. The flask should be observed u n d e r an inverted microscope daily for 48 h to ensure the absence of contamination or cytotoxicity problems that usually arise within 24 h. If these problems arise, refer to the Troubleshooting section. The flasks should then be observed every other day for the d e v e l o p m e n t of cythopathogenic effects. 7. During the incubation period, the media in the flasks is changed regularly (every 5-7 days). This is done by decanting the old media and adding fresh m e d i u m pre-warmed to 36.5 _+ I°C. The flasks are usually incubated for up to three weeks or until they develop cytopathogenic effects (CPE). The d e v e l o p m e n t of CPE is demonstrated by the extensive detachment of the cells from the monolayer or by the observation of drastic morphological changes
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8.
9.
10.
11.
12.
13.
of cells. The most COlTlrnon morphological change is the rounding u p of the cells, and their eventual lysis. CPE d e v e l o p m e n t is usually indicative of the presence of infectious viral agents. However, all CPE positive cells must be first verified before the sample is d e e m e d to contain infectious viral agents. This is done by freezing all tile CPE positive flasks at -70°C, followed by rapid thawing for two cycles of freezing and thawing. The flask contents are then filtered through a 0.2 t_ml filter, and then inoculated onto new freshly passed cell culture flasks containing confluent monolayers. Steps 3-7 are repeated. All positives must be verified at least once. In addition, all samples that do not demonstrate CPE after the three-week incubation are also f r e e z e / t h a w e d twice and passed onto new fresh flasks containing confluent monolayers and steps 3 7 are repeated. All negatives must also be verified at least once. if a negative sample comes up positive after its passage, then the positive has to be verified as in step 8. Usually injured viruses do not exhibit CPE on their first inoculation, but do so after subsequent passage. At this point the remaining subsarnples of tile water sample or viral concentrate are rapidly thawed and inoculated as mentioned above. If more than seven of the ten initially inoculated flasks (step 3) exhibit CPE then five-fold dilutions of samples are necessary. Usually, five-fold dilutions of tile sample are made in sterile 0.15 M Na~HPO,. Samples that develop CPE in the first and second passage (or ones that s h o w e d no CPE on the first but did on the second and fllird passage) are scored as confirmed positives. Samples that show no CPE on the first and second passage are scored as negatives. All CPE positive samples should be stored at 70°C for research purposes or for identification by serum neutralization tests (Melnick et al., 1973) or gene p r o b i n g . By assaying various volumes and dilutions of a sample and determining the presence of viruses by the exhibition of CPE, a most probable n u m b e r (MPN) analysis can be performed to estimate tile n u m b e r of viable infectious units in a sample. MPN calculation can be p e r f o r m e d using the folk)wing formula: MPN m l = P/(N.Q) '-~,where P equals the total n u m b e r of confirmed positive samples for all dilutions, N equals tile total v o l u m e of the assay sample (in ml) inoculated for all dilutions, and Q equals the total v o l u m e (in ml) of sample inoculated onto cultures that remained CPE negative. MPN values for the assay of undiluted samples can be confirmed with the formula: MPN = hffq/t~), where q equals tile n u m b e r of CPE negative cultures and i~ equals the total n u m b e r of cultures. MPN per liter value of the original water sample can be calculated by multiplying the MPN value by the n u m b e r of milliliters of the final concentrated sample inoculated and dividing by the total v o l u m e of water sampled.
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Troubleshooting Problems with this procedure arise from the presence of contaminating microorganisms or cytotoxic compounds in the water sample. These may also arise from the premature death of the cells on the flasks or their rounding up. The former problems usually arise within 24-48 h after inoculation while the later ones arise usually after 12-14 days. Contamination problems are evident by the presence of turbid growth in the cell culture flask's media. Cytotoxicity is marked by morphological changes of the monolayer and apparent 'unhealthiness' of the cells as compared to the negative control cells. Cell rounding up and detachment may occur during growth and mitosis. Samples should not be considered to be positive unless there are significant clusters of rounded up cells over and beyond what is observed in the uninfected controls. Photomicrographs demonstrating CPE are available in Malherbe and Strickland-Cholmley (1980). To avoid bacterial and fungal contamination problems, it is advised that the sample be first passed through a 0.22 lJm filter before inoculation onto the cells. If these problems still arise, then the media on the cells must be changed more frequently (2-3 days). This allows the antibiotics and antimycotics in the media to control the growth of the contaminants. In addition, decanting the inoculum after its incubation on the cell monolayer (step 3) also aids in reducing contamination problems but may reduce assay sensitivity. In order to reduce cytotoxicity, the inoculum from cell culture vessels after the adsorption period (step 3) is decanted and saved. A volume of 0.25 ml of pre-warmed (36.5 _+I°C) PBS or the sodium phosphate solution for each cm' of cell surface area is added into each vessel. The washing solution should be added to the side of the cell culture vessel opposite the cell monolayer. The washing solution should be gently swirled across the monolayer a minimum of two times and then decanted. Step 4 of the above procedure is then continued.
M o l e c u l a r d e t e c t i o n of enteroviral nucleic acids Principle and applications Within tile last 15 years nucleic acid hybridization and the polymerase chain reaction (PCR) have shown promise in rapidly detecting human viruses and other fastidious microbial pathogens in the environment (Atlas et al., 1992). PCR has been used extensively to detect bacterial pathogens in marine systems, particularly when target organisms are difficult to culture (i.e. viable but not culturable bacteria). Molecular detection methods have been especially important in studying the occurrence of Hepatitis A and Norwalk-like viruses, which are epidemiologically important pathogens but do not produce CPE readily in cell culture. Molecular techniques have also enhanced the speed and sensitivity of detection for the more routinely cultured enteroviruses. Given the time required to assay water samples for infectious human
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viruses by cell culture, several researchers have opted to test concentrated samples for viral RNA using reverse transcriptase-polymerase chain reaction (RT-PCR) (Griffin et al., 1999). Although this assay does not permit any differentiation between infectious and non-infectious viral particles it does allow for a fast and relatively inexpensive method to test waters for potential impact from these viruses. Furthermore, data show that enteroviruses can be detected in marine waters by RT-PCR far longer than they can be detected by cell culture (J. Jarrell et al., unpublished data) and therefore can serve as an indicator of prior sewage impact. An integrated cell culture RT-PCR method has been suggested as an appropriate compromise between these two modes of detection (Reynolds el al., 1996). In this approach a cell monolayer is inoculated (as described above), after some period of incubation (24 h to 6 weeks), the cells are frozen and thawed and the lysates are assayed by RT-PCR. This approach has proven more sensitive than strict cell culture, because some infectious viruses do not produce CPE and are, therefore, only readily detected by molecular techniques. In general, this approach reduces the possibility that non-infectious viruses will be detected by RT-PCR while still allowing a faster turn-around time because it is not necessary to wait the 6-9 weeks often required in the cell culture assay. Given the variety of primers, probes and reaction conditions already available or reported, this section will only generally describe the procedure for extracting viral nucleic acids and amplifying specific sequences using RT-PCR. There are various sources that describe the basics of PCR and RT-PCR, i.e. Innis (1990). For specific references on enteric viruses, see Griffin at al. (1999), De Leon el al. (1990), Reynolds at al. (1996), Schwab at al. (1996) and Wyn-Jones et al. (1995). Given their relatively widespread occurrence, owing partly to the Sabin (oral) polio vaccine, assays for the panenteroviral group are most commonly used. While the following highlights this protocol, the principles are the same for detecting other RNA viruses.
Materials and reagents • Thermocycler (e.g. Perkin Elmer) • Hybridization incubator • Water bath • Microcentrifuge • 1.5 ml sterile centrifuge tubes • 0.2-0.5 ml thin walled PCR reaction tubes • Milli-Q, or other ultra-purified, water • Vortex mixer • Micropipettes with appropriate tips (sterile and aerosol resistant) • RNA extraction kit (e.g. RNeasy from Qiagen), alternatively other chemicals may be used for extraction (chloroform, guanidinium isothiocyanate, etc.; Sambrook et al., 1989) • Non-charged nylon hybridization membrane (e.g. Magna GRAPH from MSI/Osmonics)
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Oligonucleotide primers and labeled (e.g. biotin, digoxygenin, 32p, etc.) oligonucleotide probe complementary to a portion of the amplified region. For enteroviruses, the most commonly used primers amplify a 197 bp section of the highly conserved 5' untranslated region (De Leon et al., 1990, Schwab et al., 1996).These primer and probe sequences are listed below: upstream primer: 5'-CCT CCG GCC CCT GAATG-3' downstream primer: 5'-ACC GGATGG CCA ATC CAA-3' internal probe: 5'-TAC TTT GGG TGT CCG TGTTTC-3' For the RT reaction, either the downstream primer or random hexamers can be used. RT and PCR reagents available in a kit form from PE Biosystems, RNAPCR Core Kit) which include reverse transcriptase (MLV), RNase inhibitor, random hexamers, 25 mM MgCI2, 10x PCR Buffer II (without added MgCI2), dNTPs and Taq polymerase.
Extraction procedures Nucleic acid m a y be extracted directly from beef extract concentrates or from inoculated cell culture flasks. M a n y methods are available for this procedure and there is little consistency in the published literature. Readers are encouraged to use any available technique for this step; however, it is imperative that the extracted material be as clean as possible because c o m p o u n d s such as humic acids inhibit the RT and PCR enzymes. The readers should also be aware that beef extract inhibits the reactions and special care should be taken to adequately remove this from the extract. In our lab, we have had success using commercially available kits, such as RNeasy from Qiagen. N e e d e d reagents and tubes are provided with these kits and directions can be followed with little or no modification. In the case of the RNeasy kits, 100 pl sample aliquots are boiled in extraction buffer to release RNA from the capsid. After a series of washes, extracted RNA is eluted from the silica-based column and suspended in 30-50 pl of RNase-free water. At this point RNase inhibitor can be a d d e d to the extracts and used directly for RT-PCR or stored for up to 1 week at -70°C (we have found that longer storage times result in some degradation and loss of signal intensity in detection steps).
Amplification procedures Briefly, PCR and RT-PCR are cyclic enzyme-based reactions that amplify target regions of DNA or cDNA. In RT-PCR, RNA is first reversed transcribed to cDNA before PCR can proceed. This step does not need to be sequence specific and a variety of 'primers' can be used for this reaction (random hexamers, oligo dTs, the PCR d o w n s t r e a m primer). In traditional PCR/RT-PCR a 100pl reaction v o l u m e is used. Concentrations of primers, reagents and sample v o l u m e should be
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optimized along with cycle parameters to yield the best results. Generally, protocols described in the manufacturer's literature (PE Biosystems RNAPCR Core Kit) offer a good starting point. Consult the owner's manual to program the thermocycler. In our lab we use the following conditions to detect enteroviruses (using previously listed primers). 1. Master Mix i reagents (RT reaction). Volumes are given per reaction/ sample for 100 121reaction. This should be prepared on ice. Reagent MgCI~ 25 mM 10x PCR Buffer II dNTPs RNase inhibitor Reverse transcriptase random hexamers (as provided)
Volume 6 t21 3 lJl 2.5 lJl each 0.9 p1 (20 U) 0.9 lal (50 U) 1.0 1M
2. The master mix should be vortexed and separated into individual 20.0 p1 aliquots to which 10 p1 of extracted sample should be added (0.2-0.5 ml thin walled reaction tubes). If the available thermocycler does not have a heated lid for the sample block, each sample should be overlaid with a drop of sterile mineral oil. Tubes should then be placed in the thermocycler, preprogrammed for 20 min at 23°C (primer annealing), 60 min at 42°C (cDNA transcription) and 5 min at 99°C to inactivate the reverse transcriptase. 3. Following the RT reaction, master mix II (PCR) should be prepared and aliquoted into each reaction tube (68 lJl per reaction). 4. Master mix II should be prepared on ice. The following volumes are for one reaction. Upstream primer Downstream primer MgCI~ 25 mM 10X PCR Buffer II Milli-Q water ~q polymerase
0.15 ~tM (-0.5 tJl) 0.15 gM (-0.5 pl) 3.6 1.d 7.2 ~I1 to final volume of 100 1-11 O.5 ~i1 (2.5 U)
. Tubes should be replaced in the thermocycler, preprogrammed for 40 cycles at 95°C, 55°C and 72°C for 1.5 min each. The samples can be stored at 4 ° C for further processing.
Visualizing amplified product After amplification by PCR, the product can be run on a 1-1.5% agarose gel and visualized by ethidium bromide staining. For the previously described procedure, the target enterovirus sequence is 197 bp. To verify the band as enterovirus, the product should be confirmed using hybridization with a labeled internal oligonulceotide probe. Again, several methods are available for this step and may vary depending on blot type (Southern transfer versus dot blot) and label used. Molecular cloning guides may be consulted for any of the techniques.
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ENTERIC PROTOZOAN
PARASITES
Background Cryptosporidium spp. and Giardia spp. are enteric protozoan parasites that commonly cause waterborne gastroenteritis. These organisms produce environmentally stable cysts (Giardia) and oocysts (Cryptosporidium) which offer protection against disinfection and environmental stresses. Relatively little is known of the prevalence and survival of Cryptosporidium and Giardia in coastal marine waters. However, studies in Hawaii have demonstrated that survival of Giardia may be closely related to salinity and declines rapidly at salinities near 35 %o. Cryptosporidium is more tolerant of these salinities; 3-4 days were required for one-log inactivation (Johnson et al., 1997). Cryptosporidium, in particular, may accumulate in certain marine environments. Fayer et al. (1997) demonstrated that the Eastern Oyster (Crassostrea vi~ginica) could effectively accumulate oocysts in the gills, hemocytes and digestive tract. Additionally, oocysts remained infective within the oyster tissue. Cryptosporidium spp. have also been isolated from the digestive tracts of Green Turtles (Chelonia mydas) (Graczyk et al., 1997). In Florida, both Cryptosporidium and Giardia have been detected in contaminated, tidally influenced streams (Phillippii Creek, Sarasota) (Lipp et al., in press).
Collection of Cryptosporidium and Giardia from m a r i n e waters Principle and applications Much like the enteric viruses, the protozoal pathogens Cryptosporidium and Giardia occur in low numbers in most marine waters and therefore must be concentrated for detection. However, the relatively large size of these organisms (N1-4 [am) permits mechanical capture with _<1 [am poresized filters. Because filters will capture both organisms, most techniques allow the simultaneous detection of Cryptosporidium and Giardia. Therefore, unless otherwise stated, the procedures described in the following sections apply to either organism. Procedures were adapted for marine water from the following sources: Standard Methods for the Examination of Water and I4~astewater (APHA 1997), Microbiological Methods for Monitoring the Environment (EPA 60018-78-017), and Information Collection Rule: Methods for Analysis of Protozoa and Viruses (US EPA 1992). The efficiency of recovery of cysts/oocysts during the processing of a sample for Cryptosporidium and Giardia have been evaluated previously by several researchers (Ongerth and Stibbs, 1987; Rose et al., 1988; 1989; LeChevallier et al., 1990; 1991a; Rose et al., 1991). The following recovery efficiencies have been reported: sample collection, 88-99%; filter elution, 16-78%; concentration and clarification, 66-77%; microscopic detection (IFA only), which also represents the overall recovery efficiency, 9-59%. Consequently, current methods for recovering and detecting protozoa underestimate the true concentrations in environmental samples. New
572
methods, including immunomagnetic separation (IMS), show promise in increasing clarification of concentrated samples and improving efficiency. Cell culture techniques under development allow for the specific detection of infectious oocysts (Jakubowski et al., 1996). These procedures will allow for a better assessment of enteric protozoa in the environment. The basic equipment used for the concentration of Cryptosporidium and Giardia can be the same as that used for enteric viruses for large volumes, and employs a 10 inch cartridge filter holder and filter (hoses, flow meter, etc.) as described below. Often virus and parasite samples are collected concurrently. In order to detect low concentrations of cysts/oocysts, large volumes of water must be passed through an appropriate filter (both membrane and cartridge filters have been used with a pore size of 1.0 btm). Unfortunately, u n w a n t e d constituents of the sampled water (e.g. particulates, precipitated minerals, etc.) are also concentrated and can interfere with subsequent procedures (Rose et al., 1989; Le Chevallier et al., 1990). Most often a y a r n - w o u n d polypropylene filter with a 1.0 btm pore size (Filterite, MD) has been used for collection of enteric protozoa. Other filter systems are available (i.e. Gelman) but have yet to be evaluated in marine wa ters.
Materials and reagents
Equipment needed for the collection of protozoa is the same as that required for enteric viruses, with the following additions: • •
I0 inch, 1.0 btm nominal porosity yarn wound filter (Filterite) I gallon disposable, sealable plastic bags (i.e. Zip Lock)
Procedure
1.
2.
3.
4.
5.
The filters are housed in a cartridge holder and the water is passed through the system by a water p u m p (see virus section). Use separate equipment for each site or chlorinate between samples. A volume meter and flow regulator are attached after the filter to indicate volumes filtered and regulate filtration flow rates (4-81 min ~), respectively. After the desired volume has been filtered (usually 50 to 400 1) or the filter clogs, place the filter immediately in a large sealable bag, double bag and label. The filter can be stored at 4°C for up to days before processing. The filter should never be frozen. After sampling, rinse non-autoclavable material such as hoses, filter housings, water meters and p u m p s with -400 1 of tap water and disinfect by exposure to 10-15 mg 1 ' of chlorine (NaOC1) for 30 min. Dechlorinate the equipment by addition of sodium thiosulfate sufficient to neutralize the remaining free chlorine (40 ml of a 10% solution per 400 1 of water). If more than one sample is being collected during the same day, equipment can be disinfected using bleach and source water (rather than tap water). Before collecting the next sample, up to
573
6.
400 1 of the water to be sampled should be pumped through the inlet hose, filter housing, and pump prior to placement of the filter in the housing. If conditions of the sample collection sites are known, it is best to move from 'clean' sites to 'dirty' sites. When possible, collection should always begin with sites least likely to have parasite contamination, so that the samples most likely to contain the greatest number of parasites are collected last.
Filter elution and sample concentration
Once a sample filter is brought into the laboratory, further processing is accomplished through elution, and reconcentration/clarification. The elution is accomplished by washing the collected material from the filter. The reconcentration/clarification is accomplished through density gradients and centrifugation or immunomagnetic separation.
Materials and reagents • • • • • • • • • • • • • • • • • • • • • • •
Razor blade or scalpel (sterile) Stainless steel tray (2) Aluminum foil Stomacher machine Stomacher bags (for washing) (4) 500 ml sterile centrifuge bottles 50 ml sterile centrifuge tubes Polystyrene pipettes (25 ml, 10 ml, 5 ml, I ml) Centrifuge (capable of 2000g) Plastic beaker (3000 ml) (3) Pasteur pipettes Glass erlenmeyers (I 000 ml, 500 ml) Flasks (2000 ml, 1000 ml, 500 ml) Kimax Brand Flask with quick-release connector Electrical pump pH meter Beakers (500 ml, 1000 ml) Magnetic stirrers and round bars Gloves Absorbent paper Kim wipes Biohazard bags Eluting solution consisting of the following reagents: Sodium dodecyl sulfate (SDS) Polyoxyethylenesorbitan monooleate (Tween 80) • Phosphate buffered saline (PBS) (10× concentration) Sodium chloride (80 g) Potassium dihydrogen phosphate (KH2PO4, 2 g)
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Disodium hydrogen phosphate anhydrous (Na~HPO, I I g), or hydrated disodium phosphate (Na~HPO,. 12H~O, 29 g) Potasium chloride (KCI, 2 g) Sodium thiosulfate (Na2S203.5H20, 100 g) Commercial bleach Antifoam A (Sigma Cat #A5758) Formaldehyde
Procedure 1.
2. 3. 4. 5. 6.
7.
Cut the filter longitudinally, tease apart and separate the fibers by hand (always use gloves). Wash the fibers in 1500 ml of eluting solution (PBS with 1c/{ Tween 80 and 1 ~7, SDS) in a large volume stomacher (Stomacher Laboratory Blender model 3500; Tekrnar, Co., Cincinnati, OH) for 5 rain. Then redistribute the fibers evenly in the bag and 'stomach' for an additional 5 min. Repeat the washing with another 1500 ml of eluting solution. Using gloves, recover the entire volume of eluate by pressing the fibers by hand and distributing in 500 ml sterile centrifuge bottles. Discard the filter fibers. Centrifuge the eluate at ~180(){~ for 20 min and aspirate off the supernatant. Combine pellets into one bottle and transfer to sterile 50 ml centrifuge tubes. Centrifuge at 1800g for 20 min and discard the supernatant. Store the pellet at 4°C and process by clarification (see below) within 48 h. (For long-term storage, resuspended the pellet in an equal volume of 3.7% formaldehyde; however this CANNOT be used for PCR detection.
Clarification using density gradients Both cysts (oocysts) and debris are recovered from the filter during the elution/washing process, using detergents and physical agitation. After concentration of the eluate by centrifugation (see above), an aliquot of the pellet, which contains cysts (oocysts) and debris, is layered onto a density gradient of Percoll and sucrose (final specific gravity: 1.10, Sigma Chemical Inc.) Centrifugation then separates the cysts (oocysts) from rnuch of the debris. This semi-purified sample is then recovered from the gradient.
Materials and reagents • • • •
Sucrose (85.58 g) Percoll (sp. gr. I. 13; Sigma Cat. #P 1644) API hydrometer Ethanol
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• • • • • •
Glycerol Centrifuge (capable of 2000g) 50 ml sterile centrifuge tubes 60 ml syringe Polystyrene pipettes (25 ml, 10 ml, 5 ml, I ml) 14 gauge, I 0-cm-long, stainless steel cannula
Procedure 1.
2. 3. 4.
Add eluting solution (see above) to 1 ml of pellet to a final volume of 20 ml in a 50 ml centrifuge tube. Using a 60 ml syringe and a 14 gauge cannula, place 30ml of percoll-sucrose (specific gravity 1.10 g ml ~) at the bottom of the tube (place the cannula at the bottom of the tube and hold vertically). Tilt the cannula slightly to allow the percoll-sucrose to underlay the sample. Centrifuge at 1300g for 20 min. After centrifugation, withdraw the top 25 ml of the sample with a 25 ml pipette. Place the collected fluid in a sterile 50 ml centrifuge tube and add 25 ml of eluting solution. Centrifuge at 1800g for 20 min. Discard the supernatant by aspiration to approximately 5 ml, leaving the clarified pellet.
Clarification using immunomagnetic separation Immunomagnetic separation (IMS) consists of attachment of the target organism to an antibody-iron bead complex, after which the sample is exposed to a magnetic field in a special test tube and rack (Dynal, Inc., Lake Success, NY). The specific organism is then concentrated through magnetic-antibody capture and the remaining supernatant is removed. The organism is normally removed from the beads using 0.1 N HC1 and the beads recaptured, while the supernatant, which contains the purified organisms, is recovered. IMS allows for both purification and concentration, and greatly improves sensitivity (Gracas et al., 1999).
Materials and reagents • • • • • • • • • •
Purified, formalin-fixed Cryptosporidium parvum oocyst stock suspension Purified formalin-fixed Giardia lamblia cyst stock suspension Sterile 15 ml centrifuge tubes Polystyrene pipettes (25 ml, 10 ml, 5 ml, I ml) Micropipets (I 000 lul, 100 IJl, I 0 lal) with appropriately sized aerosol resistant tips Dynabeads GC Combo (DYNAL Kit) Sterile microcentrifuge tubes (I.5 ml-2 ml) Flat-sided Dynal L I 0 tubes (I 25 x 16 mm) Vortex mixer Rotating (end-over-end) mixer
576
• • • •
Magnetic Particle Concentrator (DYNAL MPC-I and Dynal MPC-M) Dynal Spot-On slide Microscope slides 7 5 x 3 8 mm (Coming) and cover glasses 25 mm ~ (Coming) Monoclonal Antibodies (Waterborne, Inc, anti-Giardia, anti-
Cryptosporidium ) • •
25 mm diameter, 8 lum,White SCWP, Membrane Filters (Millipore) 25 mm diameter, 1.0 lum pore size cellulose acetate membrane filters (Sartorius)
Procedure (adapted from Dynal protocol) 1.
2. 3.
4.
5.
6.
Add the non-specific binding solutions (1 ml of buffer A and 1 ml of buffer B; Dynal) and the magnetic bead solution with the antibody to _<10 ml of concentrated sample (pellet) in a sterile 15 ml centrifuge tube. Mix tubes for 60 min on an end-over-end rotating mixer at 15 to 20 rpm (Glas-Col, Terre Haute, IN). Separate the bead-bound particles (containing Ctyptosporidiunl and Giardia present in sample) by placing the tubes into a magnetic particle concentrator (Dynal MPC-1), and gently rocking for 2 min. Remove and discard the supernatant. Remove the tube from the rack and add 0.1 ml 0.1 N HC1 and 1 ml lx SL buffer A (Dynal). This step removes the bead from target organisms Transfer the sample to a microcentrifuge tube and place into a second magnetic particle concentrator (Dynal MPC-M); gently rock for 1 rain to separate the beads from the sample. Save the supernatant, and place on a glass slide or membrane filter. Dry and stain with monoclonal antibodies tagged with fluorescein isothiocyanate (FITC) and examine microscopically as described below.
Microscopic d e t e c t i o n of parasites on m e m b r a n e
filters
Principal and applications Concentrated samples are filtered and labeled with monoclonal antibodies specific to the cyst and oocyst wall using an indirect or direct fluorescent antibody (IFA) procedure. The sample can then be examined by epifluorescence microscopy for fluorescence (brilliant apple green), shape (ovoid or spherical), size and by phase contrast or Nomarski differential interference contrast (DIC) microscopy for internal features (1-4 nuclei, axonemes and median bodies) (Le Chevallier et al., 1991a). The use of antibodies labeled with fluorescein isothiocyanate (FITC) has greatly enhanced the ability to microscopically detect Cryptosporidium and Giardia in environmental samples. Fluorescence alone, however, may
577
not be sufficient for identification. Background due to naturally fluorescing organisms and non-specific binding of the antibody may decrease accurate identification. Although false positives can be problematic, Clancy et al. (1994) found that false negatives were a bigger problem. A number of IFA systems have been developed for Cryptosporidium and Giardia. The most commonly used antibodies for environmental monitoring come from Waterborne Inc. (New Orleans, LA) and Strategic Diagnostics (Hydrofluor-Combo, Newark, DE). Although some species specificity of the antibiotics has been reported, no single antibody system can be used to identify only those species or isolates associated with human infections. In addition, the viability of the cysts (oocysts) detected in environmental samples and their ability to cause an infection cannot be determined by this method. Le Chevallier et al. (1991a; 1991b) reported that 10 to 30% of the cysts (oocysts) found were empty and without internal features. It is unclear whether this was related to sample processing or simply the result of detecting empty cysts (oocysts) in the environment. Materials and reagents
• 1,4 Diazabicyclo [2,2,2] Octane (DABCO, Sigma Cat. # D-2522) • Hoefer Manifold with stainless-steel well weights • 4'6-diamidino-2-phenylindole (for DAPI staining, 2 mg/ml DAPI in absolute methanol) • Clear fingernail polish • Microscope with 20x and 100x objectives and UV light • Pasteur pipettes • Glass Erlenmeyers (1000 ml, 500 ml) • Flasks (2000 ml, 1000 ml, 500 ml) • Kimax Brand Flask with quick-release connector • Electrical pump • pH meter • Beakers (500 ml, 1000 ml) • Magnetic stirrers and round bars • Gloves • Absorbent paper • Kim wipes • Biohazard bags Procedure
1.
2. 3. 4.
Place 25 mm cellulose nitrate or cellulose triacetate membrane filters in a Hoefer filtration manifold. Smaller membranes (13 mm in diameter) can be set up in stainless steel or polypropylene housings. Presoak filters in lx PBS. Dilute the final sample, if necessary, and filter 1-10 ml. Add the fluorescent antibodies to the filters (0.5 ml for 25 mm filters) (Strategic Diagnostics (Hydrofluor-Combo, Newark, DE))
578
5. 6. 7. 8.
9.
in the filter housing. Incubate for 30 min at room temperature. If an indirect procedure is used, the filter must first react with the specific antibody and then is washed and stained with a second fluorescent-labeled antibod}, similar to above. Wash the filters thoroughly with l x PBS (pH 7.0). Rinse the filters with a series of ethanol solutions (10~/~, 4{Y~, 6()~J< and 80'~) for dehydration and clearing. Mount the filters on a glass slide with ~60 btl of mounting media (Dabco -~ glycerol). Incubate the filters at 37°C for 20 min until the filters are clear. Add 15 to 20 btl of Dabco to the top of the filter and cover with a coverslip. Seal using clear fingernail polish. Enumerate Cryptosporidium and Giardia using a fluorescent microscope. Scan the entire filter using a 20x objective.
Identification of parasites
Procedure Initial examination of the filters is accomplished u n d e r low magnification and epifluorescent illumination. Identification of Cryptosporidium and Giardia is accomplished using 100× magnification as follows: 1. Target organisms fluoresce bright green (demonstrating monoclonaI antibody specificity). 2. Cryptosporidium and Giardia can be further distinguished by their size and shape. Giardia cysts are oval and 10 to 20 bml long by 5 to 15 bun wide. Cryptosporidium oocysts are spherical and usually 4-6 ~tm in diameter. Record the size and shape. 3. The appearance of a characteristic fold in the oocyst wall is another determinant feature for Cryptosporidium. 4. Lastly, examine suspect cysts (oocysts) for internal feature. Nucleus, axonemes or median bodies are c o m m o n l y seen in Giardia cysts and sporozoites can be observed in Cryptost~oridium oocysts.
PCR-based d e t e c t i o n o f Cryptosporidium PCR-based assays have focused primarily on Cryptosporidium, while very few protocols have been published on optimization for detection of Giardia from water (Rochelle eta/., 1997a,b). One of the first published works on detection of oocysts in water targeted 18S rRNA and reported a sensitivity of 1 to 10 oocysts (Johnson et al., 1995). In most cases 100 oocysts was the limit of sensitivity with PCR in environmental samples. C h u n g ct al. (1999), for example, was not able to detect 100 oocysts per 100 1 but did detect 1000 oocysts per 1001 using quantitative PCR. PCR has also been combined with cell culture and excystation to evaluate viability and for the detection of oocysts in oysters, mussels and cockles (GomezBautista et al., 2000; Fayer et al., 1999; Chung et al., 1999; Rochelle et al.,
579
1997a,b; Di Giovanni et al., 1999). Presently, IFA is still more sensitive than PCR for the detection of oocysts, and therefore PCR has not been proposed as a screening or primary detection method. Studies at the molecular level currently focus on methods to identify C. parvum isolates of human origin, as well as genotypes that may be of either human or animal origin (zoonotic) (Patel et al., 1998; Wiedenmann et al., 1998). Both speciation and genotyping may be used for risk assessment and characterization of contaminated waters. While no single PCR protocol for Cryptosoridium has been developed, in a recent review the majority of the researchers did have some common methodological approaches (Wiedenmann et al., 1998). M a t e r i a l s and reagents
• Dry ice or liquid nitrogen • Large beaker • Hot plate • Chelex 100 resin • I×TE (Tris-EDTA pH 8.0) • Taq D N A polymerase • Thermocycler • Primers (upstream and downstream: sequence depends on target) • PCR Buffer (PE Biosystems) • dNTPs • MgCI 2 • Milli-Q water • Sterile, aerosol resistant pipette tips • 1000 ~ul,200 ~ul and 20 ~1 pipettes • 500 ~ul thin walled reaction tubes • 1.5 ml microcentrifuge tubes Procedure
1. 2.
3.
4.
5.
Use concentrates as described above. IMS processed samples are recommended to limit inhibitors. Prepare Chelex 100 resin (Bio-Rad, Hercules, CA) equilibrated 1:1 volume-to-volume mix of resin and lx Tris-EDTA buffer (lx TE, pH 8.0). Add 100 btl of prepared Chelex resin to each 100 btl aliquot of sample (pellet). Lyse oocysts by rapid freeze/thaw. Briefly alternately expose Chelex amended samples to liquid nitrogen (freezing) and boiling water (thawing) for 1 min each (eight cycles). PCR conditions are primer specific (see Table 28.1), however, the final reaction volume can be scaled up to 100 btl, using AmpliTaq Gold DNA polymerase (Roche Molecular, Branchburg, NJ) rather than a traditional 'hot start' method. Use Nested-PCR protocols to enhance specificity to the 18S rDNA target sequence. Confirmation can be made using a labeled internal probe (isotopic or non-isotopic) (Table 28.1).
580
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ISSUES
Methods have been u n d e r d e v e l o p m e n t for approximately 40 years to detect specific waterborne pathogens, n a m e l y the viruses and protozoa. Standardized procedures have been used for drinking water and are n o w being applied to marine waters and shellfish tissues. Although
Table 28.2 Some advantages and disadvantages of methods for viruses and parasites Tool
Advantage
Disadvantage
Application
Culture for
Wide-spread use (viruses) Standardized
In some cases lack of specificity (i.e. generic for enteroviruses)
To evaluate water quality and treatment, retrospectively
Cryptosporidiunt and viruses
Results take days or weeks. Detects viable microorganisms
Microscopy for protozoa
More difficult to quantitate in some cases.
For comparison to new methods and old data bases.
Measures only a fraction of the types of microorganisms present, no ability to detect viable, but non-culturable, unless combined with PCR.
When viability is of major concern
Sample preparation can interfere with detection.
Most sensitive method readily available for protozoa.
Can be used with specific stains (monoclonal antibodies, probes, fluorogenic dyes and viability stains)
Less sensitive than culture as smaller volumes are examined.
Rapid
Microscopy can be tedious.
Use for assessment of physical removal.
Quantitative Polymerase chain reaction (PCR)
Highly specific
Measures both viable and non-viable microorganisms
Rapid Maybe inhibited by sample constituents Traditionally nonquantitative Less sensitive due to volumes processed
Excellent for presence/absence assessment (i.e. physical removal and ground water impacts). Only method available for key viruses of concern. Can be applied with cell culture techniques for rapid and specific evaluation.
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guidelines have yet to be established, the data needed to address protection of public health using risk assessment procedures for the treatment of wastewater and stormwater that m a y impact coastal water is now being collected. Microbiological methods are progressing toward the ultimate goal of detecting individual microbial pathogens per unit volume. Currently all the m e t h o d s discussed have issues associated with insufficient recovery, sensitivity (or the limit of detection) and specificity. Often these problems are related to water quality, and the v o l u m e collected, processed and examined. High turbidity, suspended solids, humic acids, and changes in p H are some of the factors that interfere with the collection of microorganisms (changing adsorption by the filter for viruses and clogging the filter limiting the a m o u n t of water that can be collected). These same materials also interfere with microscopic visualization, again limiting the v o l u m e that can be processed or examined. Such materials also interfere with the culture of viruses and bacteria. In untreated wastewater, concentrations of enteric pathogens are high e n o u g h that small volumes can be collected and examined. However, in receiving waters, larger volumes are n e e d e d and limits of detection are often quite high. There are m a n y new d e v e l o p m e n t s in microbiological methods and it is possible to s u m m a r i z e their advantages and disadvantages and application to marine waters (Table 28.2). As sewage and stormwater continue to impact our near shore coastal waters, recreational beach goers and others are at risk of exposure to harmful pathogens. Therefore, it is important that methods are d e v e l o p e d and efforts are m a d e to better understand the prevalence of enteric pathogenic microorganisms in these environments.
References APHA, AWWA, WEE (1997). Pathogenic protozoa: proposed method for Giardia and Cryptosporidium spp. In: Standard Methods .fi~r the ExamiI~atioll qfl Water a11d Wastewater, 19th edn. Supplement. Washington, DC. Atlas, R. M., Sayler, G. S., Burlage, R. S. and Bej, A. K. (1992). Molecular approaches for environmental monitoring of microorganisms. BioTeclmiques 12, 716-717. Barron, A. L., Oishevsky, C. and Cohen, M. M. (1970). Characteristics of the BGM line of cells from African green monkey kidney. Archly. Gesam. Virush~rsch. 32, 389-392. Bitton, G., Farrah, S. R., Ruskin, R. H., Butner, J. and Chou, Y. J. (1983). Survival of pafl~ogenic and indicator microorganisms in ground water. Ground Water 21, 405-410. Chung, E., Aldom, J. E., Carreno, R. A., Chagla, A. H., Kostrzynska, M., Lee, H., Palmateer, G., Trevors, J. T., Unger, S., Xu, R. and De Grandis, S. A. (1999). PCRbased quantitation of Cryptosporidium parw~m in municipal water samples. 1. Microbiol. Methods 38, 119-130. Clancy, J. L., Gollnitz, W. D. and Tabib, Z. (1994). Commercial labs: how accurate are they? J. Amer. Water Works Assoc. 86, 89-97. Craun, G. E (1984). Health aspects of groundwater pollution. In: Groundwater Pollution Microbiology (G. Bitton and C. P. Gerba, Eds), pp. 135-179. John Wiley & Sons, New York.
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Dahling, D. R. and Wright, B. A. (1986). Optimization of the BGM cell line culture and viral assay procedures for monitoring viruses in the environment. Appl. Environ. Microbiol. 51, 790-912. Dahling, D. R., Berg, G. and Berman, D. (1974). BGM, a continuous cell line more sensitive than primary rhesus and African green kidney cells for the recovery of viruses from water. Health Lab. Sci. 11, 275-282. Dahling, D. R., Safferman, R. S. and Wright, B. A. (1984). Results from a survey of cell culture practices. Environ. htt. 10, 309-313. De Leon, R., Sheih, Y.-S. C., Baric, R. S. and Sobsey, M. D. (1990). Detection of enteroviruses and hepatitis A virus in environmental samples by gene probes and polymerase chain reaction. In: Proceedings qf the Water Quality Conference, San Diego, CA. American Water Works Association, 18, pp. 833-853. Di Giovanni, G. D., Hashemi, E H., Shaw, N. J., Abrams, E A., LeChevallier, M. W. and Abbaszadegan, M. (1999). Detection of infectious Cryptosporidium parwnn oocysts in surface and filter backwash water samples by immunomagnetic separation and integrated cell culture-PCR. Appl. Environ. Microbiol. 65, 3427-3432. Fayer, R., Farley, C. A., Lewis, E. J., Trout, J. M. and Graczyk, T. K. (1997). Potential role of the Eastern Oyster, Crassostrea virginica, in the epidemiology of Cryptosporidium parvum. Appl. Environ. Microbiol. 63, 2086-2088. Fayer, R., Lewis, E. J., Trout, J. M., Graczyk, T. K., Jenkins, M. C., Higgins, J., Xiao, L. and Lal, A. A. (1999). Cryptosporidium parvum in oysters from commercial harvesting sites in the Chesapeake Bay. Emere,ing Infect. Dis. 5, 706-710. Gerba, C. P., Rose, J. B. and Singh, S. N. (1985). Waterborne gastroenteritis and viral hepatitis. CRC Crit. Rev. Environ. Contlvl 15, 213-236. Goyal, S. M. (1984). Viral pollution of the marine environment. CRC Crit. Rev. Environ. Control 14, 1-31. Gomez-Bautista, M., Ortega-Mora, L. M., Tabares, E., Lopez-Rodas, V. and Costas, E. (2000). Detection of infectious Cryptosporidium parvum oocysts in mussels (Mytilus galloprovincialis) and cockles (Cerastoderma edule). Appl. Environ. Microbiol. 66, 1866-1870. Gracas, M. Das, Pereira, C., Atwill, E.R. and Jones, T. (1999). Comparison of sensitivity of immunofluorescent microscopy to that of a combination of immunofluorescent microscopy and immunomagnetic separation for detection of Cryptosporidium parvum oocysts in adult bovine feces. Appl. Environ. Micmbiol. 65, 3236-3239. Graczyk, T. K., Balazs, G. H., Work, T., Aguirre, A. A., Ellis, D. M., Murakawa, S. K. K. and Morris, R. (1997). Ccziptosporidium sp. infections in green turtles, Chelonia mydas, as a potential source of marine waterborne oocysts in the Hawaiian Islands. Appl. Environ. Microbiol. 63, 2925-2927. Griffin, D. W., Gibson, C. J., Lipp, E. K., Riley, K., Paul, J. H. and Rose, J. B. (1999). Detection of viral pathogens by reverse transcriptase PCR and of microbial indicators by standard methods in canals of the Florida Keys. Appl. Environ. Microbiol. 65, 4118-4125. Havelaar, A. H., Olphen, M. V. and Drost, Y. C. (1993). F-specific RNA bacteriophages are adequate model organisms for enteric viruses in fresh water. Appl. Environ. Microbiol. 59, 2956-2962. lnnis, M. A. (Ed.) (1990). PCR Protocols: A Guide to Methods and Applications. Academic Press, San Diego, CA. Jakubowski, W., Boutros, S., Faber, W., Fayer, R., Ghiorse, W., LeChevallier, M., Rose, J., Schaub, S., Singh, A. and Stewart, M. (1996). Environmental methods for Cryptosporidium. J. Amer. Water Works Assoc. 88, 107-121. Johnson, D. C., Enriquez, C. E., Pepper, I. L., Davis, T. L., Gerba, C. P. and Rose, J. B.
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(1997). Survival of Giardia, Cryptosporidium, poliovirus and Salmonella in marine waters. Water Sci. Technol. 35(11-12), 261-268. Johnson, D. W., Pieniazek, N. J., Griffin, D. W., Misener, L. and Rose, J. B. (1995). Development of a PCR protocol for sensitive detection of O2~lptosporidium Oocysts in water samples. Appl. Envirolz. Microbiol. 61, 3849-3855. LaBelle, R. and Gerba, C. P. (1981). Investigations into the protective effect of estuarine sediment on virus survival. Water Res. 16, 469-478. Le Chevallier, M. W. and Trok, T. M. (1990). Comparison of the zinc sulfate and immunofluorescence techniques for detecting Giardia and Cryptosporidium. I. Amer. Water. Works Assoc. 82, 75-82. Le Chevallier, M. W., Norton, W. D. and Lee, R. G. (1991a). Occurrence of Crypto sporidilnn and Giardia spp in surface water supplies. Appl. EnvirolT. Microbiol. 57, 2617-2616. Le Chevallier, M. W., Norton, W. D. and Lee, R. G. (1991b). Giardia and Ciyptosporidium in filtered drinking water supplies. Appl. Environ. Microbiol. 57, 2617-2621. Lipp, E. K. and Rose, J. B. (1997). The role of seafood in foodborne diseases in the United States of America. Rev. Sci. Tech. Off.int. Epiz 16, 620-640. Lipp, E. K., Vincent, R., Kurz, R. C., Rodriquez-Palacios, C., Farrah, S. R. and Rose, J. B. (in press (a)). Seasonal variability and weather effects on fecal pollution and enteric pathogens in a subtropical estuary. Estuaries. Lipp, E. K., Farrah, S. R., Rose, J. B. (in press (b)). Assessment and impact of microbial fecal pollution and human enteric pathogens in a coastal community. Marine Pollution Bulletin. Malherbe, H. H. and Strickland-Cholmle~, M. (1980). Viral Cytopath~do~y. CRC Press, Boca Raton, EL. Melnick, J. L., Rennick, V., Hampil, B., Schmidt, N. J. and Ho, H. H. (1973). Lyophilized combination pools of enterovirus equine antisera: preparation and test procedures for the identification of field strains of 42 enteroviruses. Bull. W~rld Health Organizati~n 48, 263-268. Melnick, J. L. and Gerba, C. P. (1980). The ecology of enteroviruses in natural waters. CRC Crit. Rev. Environ. Control 10, 65-93. Metcalf, T. G. and Stiles, W. C. (1967). Enteroviruses within an estuarine environment. Amer. J. Epidemiol. 88, 379 391. Morris, R., and Waite, W. M. (1980). Evaluation of procedures for recovery of viruses from water II Detection systems. Water Res. 14, 795-798. Muir, P., K~immerer, U., Korn, K., Mulders, M., P6yr}; T., Weissbrich, B., Kandolf, R., Cleator, G., Van Loon, A. For the European Union Concerted Action on Virus Meningitis Encephalitis. 1998. Molecular typing of enteroviruses: current status and future requirements. Clinical Microbiolo~,,y Reviews 11, 202-227. Ongerth, J. E. and Stibbs, H. H. (1987). Identification of Cryptosporidium oocysts in river water. Appl. Environ. Microbiol. 53, 672-679. Patel, S., Pedraza-Diaz, S., McLaughlin, J. and Casemore, D. P. (1998). Molecular characterisation of Cryptosporidium parvum from two large suspected waterborne outbreaks. Commun. Dis. Public Health 1, 231-233. Paul, J. H., Rose, J. B., Jiang, S., Zhou, X., Cochran, P., Kellog, C., Kang, J. B., Griffin, D., Earrah, S. and Lukasik, J. (1997). Evidence for groundwater and surface marine water contamination by waste disposal wells in the Florida Keys. Water Res. 31, 1448-1454. Payment, P., Fortin, S. and Trudel, M. (1984). Ferric chloride flocculation for nonflocculating beef extract preparations. Appl. Environ. Microbiol. 47, 591-592. Payment, P., Trudea, M. and Plante, R. (1985). Elimination of viruses and indicator bacteria at each step of treatment during preparation of drinking water at seven water treatment plants. Appl. Environ. Microbiol. 49, 1418-1428.
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Reynolds, K. A., Gerba, C. R and Pepper, I. L. (1996). Detection of infectious enteroviruses by an integrated cell culture-PCR prodecure. Appl. Eilviroll. Microbiol. 62, 1424-1427. Rochelle, I~ A., Ferguson, D. M., Handojo, T. J., DeLeon, R., Stewart, M. H. and Wolfe, R. L. (1997a). An assay combining cell culture with reverse trnscriptase PCR to detect and determine tile infectivity of waterborne C1yptosporidium parwlm. Appl. Environ. Microbiol. 63, 2029-2037. Rochelle, P. A., DeLeon, R., Stewart, M. H. and Wolfe, R. L. (1997b). Comparison of primers and optimization of PCR conditions for detection of Oyptosporidium 19arz~llttland Gardia lamblia in water. Appl. Environ. Microbiol. 63, 2029-2037. Rochelle, P. A., Jutras, E. M., Atwill, E. R., DeLeon, R. and Stewart, M. ft. (1999). Polymorphisms in the [3-Tubulin gene of Cryptosporidium parvum differentiate between isolates based on animal host but not geographic origin. ]. Parasitol. 85, 986-989. Rose, l- B., Darbin, H. and Gerba, C. I~ (1988). Correlations of the protozoa, C~yptosporidium and Giardia with water quality variables in a watershed. Water Sci. Technol. 20, 271-276. Rose, J. B., Landeen, L. K., Riley, K. R. and Gerba, C. P. (1989). Evaluation of immunofluorescence techniques for detection of Cryptosporidium oocysts and Giardia cysts from environmental samples. Appl. Environ. Microbiol. 55, 3189-3195. Rose, J. B., Gerba, C. P. and Jakubowski, W. (1991). Survey of potable water supplies for Cryptosporidium and Giardia. Environ. Sci. Technol. 25, 1393-1400. Sambrook, J., Fritsch, E. E and Maniatis, T. (1989). Molecular Cloning, 2nd edn. Cold Spring Harbor Laboratory Press, Plainview~ NY. Schmidt, N. J., Ho, N. H., Riggs, J. L. and Lennette, E. H. (1978). Comparative sensitivity of various cell culture systems for isolation of viruses from wastewater and fecal samples. Appl. Environ. Microbiol. 36, 480--486. Schwab, K. J., De Leon, R. and Sobsey, M. D. (1996). Immunoaffinity concentration and purification of waterborne enteric viruses for detection by reverse transcriptase-PCR. Appl. Envi~viI. Microbiol. 62, 2086 2094. Unites States Environmental Protection Agency (1992). Proposed method for Giardia cysts and Cryptosporidium oocysts in low turbidity water by a fluorescent antibody procedure. D-19 Proposal P229. Unites States Environmental Protection Agency (1995). Monitoring requirements for public drinking water supplies: Proposed Rule. Federal Register 59 (28). Wiedenmann, A., Kruger, P. and Botzenhart, K. (1998). PCR detection of Cryptosporidium parvum in environmental samples--A review of published protocols and current developments. J. hzdust. Microbiol. Biotechnol. 21, 150-166. Wyn-Jones, A. P., Pallin, R., Sellwood, J. and Tougianidou, D. (1995). Use of the polymerase chain reaction for the detection of enteroviruses in river and marine recreational waters. Water, Sci. Technol. 31,337-344.
List of suppliers Dynal, Inc.
American Type Culture Collection
5 Delaware Drive New Hyde Park N Y 11042-1100, USA (800) 638-9416
12301 Parklawn Drive Rockville M D 20852, USA (800) 638-6597
IMS kits a n d e q u i p m e n t
Cell lines 586
Sartorius Corp. 131 Heartland Boulevard Edgewood NY 11717-8358, LISA (516) 254-4249
Fisher Scientific 711 Forbes Average Pittsburgh PA 15219, USA (800) 766-7000
Coming, Costar, MSI, Millepore and MediaTech Products General lab supplies (may also be ordered through any major scientific supply company) Home improvement stores (i.e. Home Depot, Lowe's)
Gasoline powered pumps, hoses, fittings and connectors, 30 gallon trash cans MediaTech Inc. P.O. Box 68 Grand Island N Y 14072, USA (800) CELLGRO
www.sartorlllS.COttl
Membranes Sigma Aldrich Co. P.O. Box 14508 St Louis MO 63178, USA (800) 325-3010
Chemicals Strategic Diagnostics 128 Sandy Drive Newark DE 19713, USA (302) 456-6789
Cell culture media
Hydrofluor Combo Kits - - FITC staining kit for Cryptosporidium and Giardia
PE Biosystems (formerly Perkin Elmer) 850 Lincoln Centre Drive Foster City CA 94404, USA (800) 345-5224
USF Filtration and Separation, Filterite Division 2118 Greenspring Drive Timonium MD 21093, USA (410) 252-0800
PCR and RT-PCR reagents
Cartridge filters and housings for viruses and protozoa
Qiagen, Inc. - USA 28159 Avenue Stanford Valencia CA 91355, USA (800) 426-8157
RNeasy Total RNA extraction kits
Waterborne Inc. 6047 Hurst Street New Orleans LA 70118, USA (504) 895-3338 www.7~atet'boFlle[llC.COt?l
Cryptosporidium oocysts and Giardia cysts
587
l 1. COLLECTION Collect water in >_100 1 container using gas driven or electric pump
2. ELUTION Force 1 I BE/G. pfl 9.0 9.5. through filter, collect eluate in sterile bottle. Neutralize pH
3. CONCENTRATION Reduce to ptf 3.5, stir and centril'uge (discard supernatant)
Reduce pit to 3.5 and add AICI3
Dissolve precipitate in NazI-IPO4
Pump conditioned water through negatively charged cartridge filter (record volume)
Adjust to pH 9,0 - 9.5 and centrifuge (save supernatant and record final volume)
4. DETECTION Inoculate cell monolayer with concentrated sample (supernatant)
Extract RNA from concentrated sample or cell culture lysates
Observe cells l\~r cytopathogenic effects (CPE) using inverted microscope
Amplify target viral nucleic acid sequence using RT-PCR Visualize results by get etcctrophoresis and internal probe hybridization
Figure 28.1. Flow chart for collection and detection of enteric viruses in marine waters.
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29 Methods for Psychrophilic Bacteria JP B o w m a n School of Agricultural Science,University of Tasmania,Hobart, Tasmania,Australia
CONTENTS
Introduction Isolation of psychrophiles Determination of cardinal temperature values Analysis of fatty acids Phenotypic characterization of marine psychrophiles
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INTRODUCTION Life forms proliferating in perpetually cold environments have developed ecophysiological and biochemical adaptations to optimize their activity at low temperatures. A broad spectrum of life is cold adapted and includes bacteria, archaea and simple and complex eukaryotes ranging from algae to fish. Organisms which are cold adapted are often referred to as psychrophiles, a w o r d derived from ancient Greek and Latin meaning literally cold-loving (psychros cold, philus/phile lover, loving). In the e p o n y m o u s review on bacterial psychrophiles by Morita (1975), a 'true' psychrophile included any organism able to grow optimally at about 15-20°C or less but was unable to grow at room temperature (20-25°C) or higher. Research suggests true psychrophiles are comparatively u n c o m m o n as the majority of bacteria isolated in cold ecosystems are what can be termed psychrotrophic or facultative psychrophiles, or m u c h more appropriately psychrotolerant (Nichols et al., 1995). These bacteria have temperature optima at 20°C or more but are able to grow at 0°C. Indeed, the growth rates of psychrotolerant bacteria are usually equivalent to or better than that of psychrophiles at low temperatures. Psychrotolerant bacteria a b o u n d even in the coldest of environments, simply because m a n y of them are ecophysiologically resilient and nutritionally versatile species. The aim of this chapter is to provide an overview of various procedures useful for the isolation and study of a specific group of bacteria, the psychrophiles. Most procedures detailed are general m e t h o d s modified for cold-adapted bacteria but are broadly applicable in the s t u d y of
METHODS IN MICROBIOLOGY,VOLUME30 1SBN0-12-521530 4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any form reserved
marine bacteria. The chapter first covers isolation, routine cultivation and maintenance of psychrophiles. Procedures to accurately determine cardinal growth temperatures using temperature gradient incubator (TGI) are then detailed. Techniques for definitively identifying and quantifying fatty acids implicated in cold adaptation of cellular membranes including polyunsaturated fatty acids (PUFA) and branched chain fatty acids are explained. Phenotypic characterization for identification and taxonomic purposes is also covered including a list of phenotypic tests applicable to studying marine psychrophiles and marine bacteria in general.
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ISOLATION
OF PSYCHROPHILES
Principle and applications Psychrophilic taxa are found only in permanently cold habitats, environments which have constant annual temperatures of less than 4°C (Morita, 1975) and which are not affected by periodic or intermittent solar insolation. The marine ecosystem appears to be an excellent place to find psychrophilic bacteria, as most of the oceanic volume is cold (<5°C). Marine psychrophiles almost without exception are able to grow at temperatures down to the limit at which normal seawater stays liquid (-2 to -5°C). Most known psychrophiles originate from the marine environment, isolated from deep waters, sea-ice and sediment (Table 29.1). Some anecdotal evidence also suggests ideal environments for isolating psychrophiles are both permanently cold and relatively eutrophic. For example Burton Lake, a eutrophic marine basin located in Eastern Antarctic, has enriched psychrophilic bacterial populations throughout its waters (Nichols et al., 1995) while adjacent coastal waters have only low psychrophilic bacterial populations (Delille, 1996). Psychrophilic populations, like other marine heterotrophic bacteria, are closely coupled to primary production (Helmke and Weyland, 1995). Psychrophiles may also abound in attached communities on marine snow and organic detritus or on the surfaces of marine fauna and flora. Data from sea-ice communities suggest species of the order Cytophay(ales and gamma proteobacteria make up the majority of marine psychrophiles (Bowman et al., 1997). This has also been borne out in 16S rDNA base cloning library analyses (Brown and Bowman, unpublished). Species of the order Cytophagales appear to be closely associated with phytoplankton while gamma proteobacteria are more likely to be free-living or associated with marine fauna. Relatively few psychrophiles so far have been isolated from the oceanic pelagic zone but isolates from the deep ocean are usually barophilic or barotolerant and specialized methods may be required for their study (described in detail in Chapter 30 of this volume). Overall, there still appears to be huge scope for the isolation of novel psychrophiles from the marine environment. A small number of psychrophilic species have been isolated from the terrestrial environment. 592
There are no specific protocols, such as specific media or enrichment strategies available for the isolation of psychrophiles. Though the methods described below concentrate on the isolation of psychrophilic chemoheterotrophic bacteria, theoretically prokaryotes of any functional group have psychrophilic equivalents, for example psychrophilic methanogens, sulfate reducers and methanotrophs have recently been described (Table 29.1). Thus, existing isolation protocols for bacteria with distinct functionalities can be adapted by applying the general approach described here.
Samples Seawater, seawater suspensions of marine sediment, suspensions of detritus and marine snow and pieces/swabs of the surfaces of marine fauna and flora samples can be added directly to or spread/placed onto isolation media and incubated at temperatures of 0-4°C. The samples to be investigated should be stored at all times at 0-4°C until used in experiments; however, brief exposure to higher temperatures (<25°C for a few hours) will usually have only a limited deleterious effect on psychrophilic populations present. Sample should never be frozen, as many psychrophiles are very easily lysed by freeze-thawing in the absence of cryoprotectants. Incubation at very low temperatures (<0°C) has little if any benefit as psychrotolerant bacteria can grow just as fast as psychrophiles. For sea ice samples, the sea ice must be thawed in seawater at 0-4°C to prevent hypotonic shocking of the bacteria present. Incubation times are dependent on the initial populations present but in most cases, growth should be visible within 7-28 days.
Media The media useful for isolating chemoheterotrophic marine psychrophiles are shown below. All media can be solidified using agar (at 1.5%) and are sterilized by standard autoclaving (121°C, 15-20 min). The best general media for marine psychrophiles is marine 2216 agar (Difco Laboratories), alternatively it can be prepared from separate constituents in artificial seawater (ASW, see formula below) or natural seawater. If natural seawater is used for making media, ideally it should be filtered (0.21Jm pore-sized filters, Millipore) rather than autoclaved. Reports suggest better recoveries of bacteria are possible on dilute media prepared with raw filtered seawater. Dilute media, including the SWC and SWCm media of Irgens et al. (1989) and R2A medium (Oxoid) prepared in seawater, are also effective for isolating a diverse range of marine chemoheterotrophic bacteria including several difficult psychrophilic species (e.g. Colwellia, Polatvmouas and Polaribacter spp.). Media should be kept refrigerated before being used for isolation, purification or subculturing. Likewise any significant exposure of psychrophiles to temperatures greater than 25°C (more than 1-2 h) should be avoided to prevent loss of viability.
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Marine medium formula: 5 g peptone, 2 g yeast extract and I 0 mg ferric phos-
phate dissolved in 1000 ml ASW. The pH is adjusted to approximately 7.5. The formula is based on marine 2216 agar produced by Difco Laboratories. SWC medium formula (Irgens et al., 1989): 0.5 g tryptone, 0.5 g yeast extract, 0.2 g beef extract, 0.2 g sodium acetate dissolved in 500 ml of ASW and 500 ml water. Adjust pH to 7.6. SWCm medium formula (Irgens et aL, 1989): 0. I g KH2PO 4,0.001 g ferric citrate, 0.4g NH4CI, 0.4g yeast extract, 0.4g beef extract, 0.4g tryptone, 2ml Humer's mineral salts (see below) and 0.2g carbon source (optional) dissolved in I000 ml of ASW. Adjust pH to 7.6. Vitamin solution (see below) may also be added if desired at I 0 ml I ' R2A-seawater agar formula: 0.5 g yeast extract, 0.25 g tryptone, 0.25 g peptone, 0.5g casein hydrolysate, 0.5g D-glucose, 0.5 g soluble starch, 0.024g MgSO4.7H20 and 0.3 g sodium pyruvate dissolved in 1000 ml ofASW.The pH is adjusted to about 7.5. The formula based on R2A agar produced by Oxoid. Artificial seawater (ASW) formula (ZoBell, 1946): 0.002g NH~NO3, 0.027g H3BO3, I. 14 g CaCI2.2H20, 0.001 g Fe(PO4)3, 5.143 g MgCI~, 0. I g KBr, 0.69 g KCI, 0.2 g NaHCO3, 24.32 g NaCI, 0.003 g NaE 0.002 g Na203Si.9H~O, 4.06 g Na~SO, and 0.026 g SrCI~.6H20 dissolved in I000 ml of distilled water. The chemicals can be added together dry and mixed thoroughly to make a large supply (add 35 g per liter of media). Sea salts can also be purchased directly from the Sigma-Aldrich Chemical Co. Hutner's mineral salts solution (Cohen-Bazire et al., 1957): Dissolve 10g of nitriloacetic acid in 950 ml of water and neutralize by adding 7.3 g of KOH. Then add 14.45 g MgSO~, CaCl2.2H20, (NH4)6MoTO~.4H~O, FeSO4.7H20 and 50 ml stock salts solution and adjust the final pH to 6.8.The stock salts solution consists of 2.5 g ethylenediaminetetraacetic acid, 10.95 g ZnSO,.7H~O, 5.0g, FeSO,.7H20, 1.54g MnSQ.H20, 0.392g CuSO~.5H~O, 0.248g Co(NO~)~.6H20 and 0.177 g Na2B,O 7. I 0H~O in 1000 ml of distilled water slightly acidified with a few drops of sulphuric acid to prevent precipitation. Vitamin solution (Balch et al., 1979): Dissolve the following vitamins one at a time in 1000 ml of distilled water and adjust pH to 7.0 using NaOH: 5 mg p-aminobenzoic acid, 2 mg folic acid, 2 mg biotin, 5 mg nicotinic acid, 5 mg calcium pantothenate, 5 mg riboflavin, 5 mg thiamine.HCI, I 0 mg pyridoxine, 0.1 mg cyanocobalamin, 5 mg thioctic acid. Store refrigerated in the dark. Filter-sterilize before use or for long-term storage.
Enrichment and enumeration
No direct isolation methods for psychrophilic bacteria are available, however, an initial enrichment of a sample in liquid isolation media at 0-2°C appears to slightly enhance the isolation of psychrophiles (Morita, 1975). The enrichment should be carried out for up to only 24-48 h and the sample immediately plated or transferred to fresh media. The enumeration of psychrophiles is only practical where their populations are greater than that of psychrotolerant bacteria, e.g. in sea ice algal assemblages (Bowman et al., 1998d). By determining most probable number (MPN) 596
counts (see Koch, 1994 for a detailed protocol for MPN analysis) at 0-2°C and at 25°C for a given sample, the proportion of psychrophilic versus psychrotolerant bacteria can be revealed.
Routine maintenance and preservation of psychrophilic cultures General storage on agar
Many psychrophiles can be maintained on agar plates or slants for long periods at 1-2°C in a frost-free cooler or refrigerator. Care must be taken to avoid freezing of the media due to ice nucleation events as it will result in complete loss of viability of the cultures. Media should be supplemented with suitable antifungal agents such as cycloheximide (at 100 btg ml ', add from a filter-sterilized 10% ethanol stock) a n d / o r nystatin (at 250 U m l ' , add from a 25 000 U m l ' filter-sterilized methanol stock). If agar plates are used, they need to be quite dry to avoid bacterial contamination. Most psychrophiles which have been isolated should be subcultured every 4-6 months when stored at 2°C. Storage at higher temperatures (4 to 10°C) requires more frequent transfer (once every 1-3 months) as viability is lost at a higher rate. Some psychrophiles, such as the species Colwellia psych~vrythreae and some members of the Cytophagales, are quite delicate and will die on plates in only a few days. Cryopreservation
For longer term storage, make a dense suspension of cells in about 2-5 ml of growth media which has been supplemented with 20% glycerol or 20~7~ dimethyl sulfoxide. The suspensions should then be frozen initially at -20°C and then stored at -70 to -80°C. For continued recovery of cells from the frozen suspension, repeated thawing should be kept to a minimum. For most psychrophiles, inoculation of frozen culture directly to plates or liquid media is usually sufficient. Large numbers of small aliquots of the cryopreserved culture(s) m a y also be a convenient safeguard as they are thawed only once, used and then discarded. Special tubes and boxes for cryopreservation storage are available from a number of laboratory suppliers.
D E T E R M I N A T I O N OF C A R D I N A L T E M PE R A T U RE V A L U ES Principle and applications The square root growth model (Ratkowsky rt al., 1983) has been implemented to accurately determine cardinal growth temperatures of a variety of psychrophilic bacteria (Nichols and Russell, 1996; Bowman et al., 1998a,b,c). This model is based on the principal that the square root of the growth rate is linearly related to temperature and can predict growth rates
597
across the entire biokinetic range (Ratkowsky et al., 1983). The model can be defined as follows: ~, r = b ( T - TMI~,, )(1 - e ~cr-r.,~.,~/)
where r is the growth rate at temperature T, T.,~,~is the notional m i n i m u m growth temperature (where 5'r = 0), T,>x is the notional m a x i m u m growth temperature (where x/r = 0), b is the slope of the regression line and c is the coefficient to be estimated experimentally. Together with the optimal growth temperature (Topr), T~,~ and T~Ax are cardinal temperatures for the biokinetic range of a given organism. All cardinal temperatures occur over a c o n t i n u u m range including T,,,\,. For marine psychrophilic and psychrotolerant bacteria, TM,,, values are usually in the range of -5°C to -22°C. Temperature gradient incubator (TGI)-based analysis of cardinal temperatures provides a useful set of autoecological data for psychrophilic bacteria and can be used for physiological and environmental comparisons and also provides useful data for strain characterization.
Equipment and reagents •
•
• •
150 ml flasks, L-tubes with metal or plastic caps (Bellco). L-tubes are customaltered test tubes made of optical quality glass (original dimensions 25 cm long, 15 mm diameter) which have modified by glass-blowing to form an Lshape ( 18 cm stem, 7 cm side arm) which allows mixing of the cell suspension in the TGI without spillage. TGI or several waterbaths/incubators.The TGI should ideally be placed in a room that has a constant air temperature reducing the fluctuation of temperatures within the tubes. Spectrophotometer which can read glass tubes (diameter 0-15mm), e.g. Spectronic 20D. Electronic hand-held thermometer with thermocouple.
Assay 1. Inocula should be grown to the late logarithmic or stationary growth phase in a suitable growth m e d i u m and at a temperature that ensures rapid growth. For m a n y marine chemoheterotrophic psychrophiles, marine 2216 liquid broth and a temperature of about 10°C is quite adequate. 2. L-tubes containing 10 ml of the growth m e d i u m are placed in the TG! for at least 1 h to allow for temperature equilibration. The TGI should be set with a m i n i m u m temperature of about 0°C and a m a x i m u m temperature of 30°C (for psychrophiles); for psychrotolerant bacteria the m a x i m u m temperature end of the TGI should be set to about 45-50°C.
598
3. L-tubes are inoculated with sufficient growth to achieve an absorbance at 540 n m of about 0.1 (if the initial cell concentration is about 10 ~'~cells ml ' the a m o u n t w o u l d be 200-300 ~tl). 4. Optical density readings are then taken just following inoculation. 5. L-tubes are agitated at about 40 oscillations per minute to avoid formation of oxygen gradients. 6. At periodic intervals after inoculation, optical density values at 540 n m and the time since inoculation are recorded. Between optical densities of 0.01 to 1.5 turbidity increases linearly (Dalgaard et al., 1994). 7. G r o w t h is complete once optical density at 540 n m exceeds about 1.5. Only the exponential growth rate area of the growth curve is needed and recordings into the stationary growth phase are not necessary. A m i n i m u m of 15 readings should be recorded for each tube. 8. At approximately one generation intervals and following growth cessation, the temperature is recorded using an electronic therm o m e t e r fitted with a thermocouple from each L-tube. 9. Optical density versus time should form a sigmoid curve from which m a x i m u m specific growth rate (#,,,,,,) and doubling time (t,) can be d e t e r m i n e d from the steepest tangent to the fitted curve as follows: In 2 _ ln(lo)B d nlax
--
td
C1
0.08-
0.07 0.06 -
/~/~
0.05-
0.04-
0.03 0.02
-
0,01
-
0.00-
-25
-20
15
10
-5
0
5
10
15
20
25
30
Temperature (°C) Figure 29.1. Square root growth rate-temperature plots of (a) Shewanella gelidimarina and (b) Glaciecola punicea showing cardinal growth temperatures T...... T,,,,~ and T,~x. The plotlines are non-linear regressions fitted using the Macintosh program UltraFit (v 3.0). Data was adapted from Nichols and Russell (1996) and Nichols et al. (1999). 599
where B is the slope of the steepest tangent. The doubling time can be thus determined by fitting the linear section of the growth curve (exponential growth region) with a regression line and determining the time interval (in minutes) required for the optical density to double in this region. 10. To determine cardinal temperatures, the reciprocal square roots of either growth rates or generation times are taken and plotted against their respective temperatures. A non-linear regression is then fitted using appropriate software (e.g. SigmaPlot, UltraFit) (Figure 29.1).
Potential problems and limitations Growth yields at the supra- and sub-optimal temperature extremes decrease markedly. This has the effect of potentially skewing growth rate information. Thus, T ~ and T ~ values are subject to some level of error. To counter this viable count data, performed by serially diluting cultures and plating onto agar (incubated at the T~,pT)can be used to help pinpoint the temperature growth limits.
41,e,e, Hl, l, A N A L Y S I S OF F A T T Y A C I D S Principle and applications The ability to maintain cellular membranes in a homeoviscous state is an important adaptation of psychrophilic bacteria (Nichols et al., 1995). In this respect psychrophilic bacteria often express high levels of anteisoand iso-branched fatty acids and unsaturated fatty acids, depending on the taxonomic group (Nichols et al., 1995). Several psychrophiles have the ability to form polyunsaturated fatty acid (PUFA) a trait unusual among bacteria (Nichols et al., 1995; Russell and Nichols, 1999). PUFAs produced by psychrophiles include eicosapentaenoic acid (EPA, 20:5m3), docosahexaenoic acid (DHA, 22:6m3) and arachidonic acid (AA, 20:4c06), fatty acids which are important 'nutriceuticals' (Nichols et al., 1999). In this section methods for the analysis and identification of fatty acids including PUFA are given. Fatty acid analysis initially uses a modified Bligh and Dyer procedure (Bligh and Dyer, 1959; White et al., 1979) to obtain an extract of whole cell fatty acids and of neutral lipids (hydrocarbons, sterols, waxes, etc.). Fatty acids are then transesterified to methyl esters (fatty acid methyl esters, FAME) and analysed by GC-MS techniques. GConly systems which allow for rapid fatty acid analysis, such as the MID1 system, identify FAME components by retention times alone, and are not able to definitively identify all fatty acids including many mono-unsaturated fatty acids and unusual fatty acids such as PUFAs. Thus, a high proportion of the fatty acid profile can be left unidentified or even misidentified. Using GC-MS accurate identification of fatty acids can be achieved. Identification of the position of double bonds in
600
mono-unsaturated fatty acid FAME is possible by dimethyldisulphide (DMDS) derivatization (Dunkleblum et al., 1985). In this method, DMDS in a chemical reaction catalysed by iodine attacks the fatty acid at the double bond resulting in CH~S adducts which can be identified by GCMS (Figure 29.2). For more complex PUFAs such as EPA, DHA and AA the number of mass fragments derived from DMDS derivitization makes mass spectra too complicated to be interpreted, instead, PUFA FAME can be reacted with 2-amino-2-methylpropano] to create 2alkenyl-4,4-dimethyloxazoline (DMOX) derivatives (Fay and Richli, 1991) (Figure 29.2). DMOX derivatives have the advantages of having high volatility allowing direct GC analysis and their mass spectra are easily recognizable allowing unambiguous determination of the positions of unsaturation.
(a)
, SCH 3 O '2~ / II CH3(CH2)mCH=CH(CH2)nCH2COCH 3 ~ CH3(CH2)mCH--CH(CHo)nCH,:,COCH q DMDS I ~ ~ FAME SCH3 MSD
SCH3t ~ CH3(CH2)mCH--(~H(CH2)nCH2COCH 3 ~
~ CH3 CH3(CH2)mCH
I SCH3 (M')
(A')
(~ -CH3OH ~H(CH2)nGH2COCH3 ----t~ ~H(CH2)nCH=G==O SCH 3
(b)
(B')
SCH3
(C')
RC/N./"O H2NX'~ 180o(3 z ~ ' O ~ OHCH3+ HO/J ' ~ R C N o , ) " FAME
2-amino-2methytpropanol
DMOX derivative
Figure 29.2. Chemical reaction schemes for fatty acid methyl ester (FAME) derivitization for determination of double bond position. (a) DMDS derivitization showing formation of four diagnostic ions for a straight chain FAME (adapted from Dunkleblum et al., 1985); (b) DMOX derivitization (adapted from Fay and Richli, 1991).
Equipment and reagents
Lipid extraction and saponification • Freeze drying unit (optional), fume hood • Sepatory funnels and stand, GC vials with Teflon-lined screwcaps or septa • Waterbath or incubator set to 80°C • Milli-Q water or double distilled water, nanograde methanol, nanograde chloroform, nanograde hexane, potassium hydroxide, nonadecanoate (C,9 standard) • Nitrogen gas and manifold for drying and concentrating samples
601
DMDS derivitization •
Dimethyl disulphide, sublimed iodine, diethyl ether, nanograde hexane, sodium thiosulphate • GC vials with Teflon-lined screw caps • Incubator or waterbath set at 40°C • Nitrogen gas and gas manifold for concentrating samples
D M O X method
• • •
2-amino-2-methylpropanol (free base), dichloromethane, nanograde hexane, sodium sulphate GC vials with Teflon-lined screw caps Oven set at 180°C Nitrogen gas and gas manifold for drying samples
Chemicals, reagents, solvents, GC vials and other minor e q u i p m e n t can be purchased from a variety of companies including Sigma-Aldrich, Mallinckrodt, Alltech, etc. (see List of suppliers for details).
Gas chromatography-Mass spectrometry The gas chromatograph utilized should be connected to a Mass Selective Detector (MSD) (Hewlett-Packard, a m o n g various companies, produce excellent GC-MS equipment). The GC conditions given in the Assay section have been optimized for a 50 m x 0.32 m m internal diameter crosslinked methyl silicone (0.171am film thickness) fused silica non-polar capillary column. A polar phase capillary column (using the same GC conditions) can also be used to identify co-eluting components. The carrier gas used is helium and the injector and the detector are maintained at 290°C and 310°C, respectively. Operating conditions for the MSD include: electron multiplier set at 2000-2200 V; transfer line set at 300°C; autotune file DFTPP normalized; electron impact energy set at 70 eV; scan threshold set at 1500; scan rate set at 0.8 s '; and mass range to be analysed set at 40-600 atomic mass units (amu).
Assay Bligh and Dyer extraction and saponification 1. Lyophilized cells (about 10-50 mg) are weighed and a d d e d to 8 ml of water, 20 ml of a n h y d r o u s methanol and 10 ml of chloroform in a separatory funnel. The mixture is then shaken vigorously and allowed to extract for at least 6 h. Freeze-dried cells are ideal if quantification is an important issue, otherwise a cell pellet (washed twice with seawater) can also be used. 2. An additional 10 ml of chloroform and 10 ml of water is a d d e d to the suspension, mixed and phases allowed to separate. 3. The lower chloroform phase is then decanted into a round bottom flask.
602
4. Solvents are then r e m o v e d in vacuo using a rotary evaporator, and redissolved in a small v o l u m e of chloroform. The concentrated lipid extract can be stored at -20°C u n d e r nitrogen. 5. For saponification all of the chloroform is removed by using a stream of N~ and the residue is resuspended in 3 ml of 5c7~ ( w / v ) KOH in 80:20 ( v / v ) methanol:water. The mixture is then incubated for 3 h at 80°C. 6. After cooling, 1 ml of water and 1.5 ml of 1:1 hexane:chloroform are added, and the mixture is shaken vigorously. The suspension is then centrifuged and the organic phase transferred to a clean tube. The extraction is repeated twice. This yields non-saponifiable neutral lipids such as sterols, hydrocarbons and waxes. 7. To the remaining aqueous phase (containing free fatty acids), 0.5 ml of concentrated HC1 and 1 ml of water are added. This is extracted three times with 1.5 ml of hexane:chloroform as shown in step 6. The solvent is evaporated u n d e r N~. 8. The residue is dissolved in transesterification reagent (10:1:1 methanol:chloroform:HC1) and heated at 80°C for 1 h. 9. To the cooled mixture add 1 ml of water and extract three times with 1.5 ml of hexane:chloroform to yield FAME. 10. The solvents are removed by using a stream of nitrogen and the residue re-dissolved in hexane. A k n o w n a m o u n t of C,~, (nonadecanoate) or similar internal standard can then be added. The FAME is n o w ready for GC-MS analysis and can be used for DMDS derivitization and the DMOX method. D M D S derivitization
1. Samples (FAME) in 20-50btl hexane are treated with 70-100~1 of DMDS and one drop of iodine reagent (60 m g of iodine in 1 ml diethyl ether) and incubated at 40°C for 24 h. 2. Reaction mixtures are then cooled and diluted with about 200 btl of hexane. 3. The iodine is r e m o v e d by adding 100 ul of 5% aqueous sodium thiosulphate with shaking. 4. The organic phase is r e m o v e d and the aqueous phase re-extracted with 100 ~1 of hexane. 5. The extract is then concentrated to a small v o l u m e u n d e r a stream of N~. The sample is n o w ready for GC-MS analysis. D M O X method 1. Samples (FAME) are dissolved in 500 btl of 2-amino-2-methylpropanol
and heated at 180°C overnight. After the reaction mix is cooled, 5 ml of dichloromethane is a d d e d and mixed thoroughly. The mixture is then washed twice by extracting with 2 ml of distilled water. . The organic phase is then dried by adding sodium sulphate and then evaporated u n d e r a stream of N2 at room temperature. The residue is dissolved in a small a m o u n t of hexane and is ready for GC-MS analysis.
2.
603
Gas chromatograph-mass spectrometry analysis 1. Samples are injected into the GC at 50°C in the splitless m o d e with a 2 min venting time. The GC oven is p r o g r a m m e d to increase in temperature from 50°C to 150°C at 30°C min ', then at 2°C min ~until 250°C is reached and then at 1°C m i n ' until a final temperature of 300°C is attained which is maintained isothermally for 15 min. MS acquisition should be started after 7 min for FAME and about 10 rain for D M D S / D M O X adducts. Resultant chromatograms and mass spectra are then compared using appropriate software. 2. C o m p o u n d s are quantified and identified by comparison of relative retention data, peak area (in relation to the internal standard) and mass spectra with other previously reported compounds. 3. For DMDS adducts derived from m o n o - u n s a t u r a t e d fatty acids four diagnostic ions (M', A', B', C') (Figure 29.1, Table 29.2) occur while only three form for mono-unsaturated fatty alcohols and fatty aldehydes. In the case of DMOX adducts the presence of a double bond is indicated by an interruption of the regular pattern produced by successive chain cleavages of methylene units. In other words, the mass spectra will show a series of fragment clusters that are separated normally by 14 amu. When a double bond occurs the interval becomes only 12 amu between two fragment peaks. The successive fragment peaks contain n and 11-1 carbon atoms of the acid moiety and thus the double bond occurs between carbons n and n+l in the fatty acid (Fay and Richli, 1991) (Figure 29.3).
Table 29.2 Mass spectrometric data of DMDS derivatives of various FAME ~ DMDS derivatized diagnostic ions
FAME
12:1o)5 12:1o)3 13:1o)5 14:1o)9 14:1c07 14:1o)5 14:1o)3 16:1~09 16:1o)7 16:1o)5 16:1o)4 18:1o)9 18:1(o7
M+
A+
B+
C+
306 306 320 334 334 334 334 362 362 362 362 390 390
117 89 117 173 145 117 89 173 145 117 103 173 145
189 217 203 161 189 217 245 189 217 245 259 217 245
157 185 171 129 157 185 213 157 185 213 227 185 213
'Adapted from Dunkleblum et a l (1985).
604
100
IH3
90, 8070. 60. 50. m
152
4o-:
192
233
,
,od 178
ii 8t , ,t
o
5(?
1 O0
1 50
27
250 Atomic mass units 200
33
lr
12,6 2 / /~ , 4I ,L/ ~~, L.~.~. / ~]i ,a, , . .~ .. 300
350
400
Figure 29.3. Mass spectrum of docosahexaenoic acid (22:6) DMOX derivative. Double bonds (A) positions are indicated by pairs of diagnostic ions (re~z) separated by 12 amu: A4 (m/z 139), A7 (m/z 166 178), A10 (m/z 206 218), A13 (m/z 246-258), A16 (m/z 286-298) and A19 (m/z 326-338). Mass spectrum and data adapted from Fay and Richli (1991). The abundance for ions of 178 amu and greater has been increased by five times to make the mass peaks more obvious.
PHENOTYPIC CHARACTERIZATION MARINE PSYCHROPHILES
OF
P r i n c i p l e and a p p l i c a t i o n s
u ee~ oL ¢, U >,'r"
o m m ° m
This section provides descriptions for a variety of phenotypic tests useful for the characterization of psychrophilic marine bacteria. The tests are also widely applicable to other aerobic chemoheterotrophic bacteria simply by altering basal media formulation and incubation conditions. Characterization data has a variety of uses, first the data is required for taxonomic analysis, particularly if the objective is to place strains into a novel taxonomic group. In this respect other techniques are also required - - the so called polyphasic taxonomic approach - - in which phenotypic, chemotaxonomic and genotypic data are collectively analysed. Methods for c h e m o t a x o n o m y including fatty acid, phospholipid, quinone and cell wall analysis are described in detail in the literature. Genotypic analysis including DNA base composition analysis, D N A : D N A hybridization and 16S rDNA sequence analysis (see Chapter 18 of this volume) are also covered in detail in the literature and the m e t h o d s therein are broadly applicable to a l l prokaryotes. In this section, tests covered are proven useful for the characterization of marine chemoheterotrophic bacteria. However, phenotypic tests more specific
605
a-~o
"O o¢4-1 I:
for autotrophic and strictly anaerobic bacteria are not shown as very few or no psychrophiles of these physiological types have been isolated so far. Various literature sources can be consulted for tests applied to characterize recently described psychrophilic methanogens, sulphate reducers and methanotrophs. Some of the tests are applicable for screening for cold-adapted enzymes. Methods for quantitative analysis of coldadapted enzymes are much the same as for normal enzymes. The characterization, mechanistic characteristics and features of cold-adapted enzymes have been covered recently in n u m e r o u s reviews (Feller et al., 1996; Feller and Gerday, 1997).
Ecophysiological tests
Temperature range For large numbers of strains temperature optima and maxima can be roughly estimated using either liquid or solid media. More accurate indications of T;~.~., T~,,., and T,,:, x should be obtained using liquid media as there is an approximate 5°C differential in To,,, values between solid and liquid media (Bowman et al., 1998a) possibly due to the effect of desiccation. For example, Psychroflexus torquis and most Polaribacter spp. show growth up to T,~,~ values of 15-20°C in liquid media but will not grow on agar plates at temperatures above 10-12°C. Thus underestimation of optimal temperatures can occur if agar media is used. Experiments should be performed at 5°C intervals starting at 0 to -5°C.
Salinity range For marine bacteria the range of salinity at which growth occurs can be easily tested on agar plates or in liquid media and should be tested at approximately their growth temperature optima. Media lacking any a d d e d NaC1 (a very low level of Na may be derived from the organic constituents but levels are usually <1 ppt) can be adapted from the media given in the isolation section. Tests should be performed to see whether strains grow well in the absence of divalent cations, Mg e* and Ca >. Usually there is a strong difference, either the strain grows well or it does not grow at all. For strains requiring divalent cations the media can be s u p p l e m e n t e d with 50 mm MgCI: and 8 mM CaC12.2H20. As k n o w n psychrophiles appear to be almost exclusively slightly halophilic the range of salinity tests does not need to exceed a m a x i m u m of 1.5-2 M NaCI.
pH range This data is usually not necessary w h e n marine bacteria are the focus of study as without exception all have optima approaching that of seawater (pH 7-8) and have a plateau of p H tolerance ranging from at least p H 6.0 to p H 8.0. 606
Biochemical tests
Rapid tests kits can be used to determine the biochemical properties of selected strains with API 20E, API 32 AN ID and API-ZYM test strips (BioMerieux) being particularly useful. Though the strips are not designed for identification of marine bacteria, they contain a wide range of tests that do not require special media or specific conditions for use. They are also rapid and convenient though on tile other hand they are expensive. The API 20E test strip has the following tests which can be of use in characterizing marine bacteria: nitrate/nitrite reduction, arginine dihydrolase, lysine and ornithine decarboxylase, gelatin hydrolysis, indole production, H~S production from L-cysteine and tryptophan deaminase. The acid production tests of the API 20E strips are less useful as marine bacteria mav not acidify carbohydrates strongly enough to give definitive results (see below). The AP132A kit contains a large number of potentially useful enzymatic tests including: arginine dihydrolase, o~-galactosidase, [3-galactosidase, [3-galacto(6-phospha te)sidase, cz-glucosidase, [3-glucosidase, o~-arabinosidase, 13-glucuronidase, ~-N-acetylglucosaminidase, R-fucosidase, alkaline phosphatase, glutamate decarboxylase, urease, arginine arylamidase, proline arylamidase, leucyl glycine arylamidase, phenylalanine arylamidase, leucine arylamidase, pyroglutamate arylamidase, tyrosine arylamidase, alanine arylamidase, glycine arylamidase, histidine arylamidase, glutamyl glutamate arylamidase, serine arylamidase, mannose and raffinose fermentation, nitrate reduction and indole production tests. Many of the same arylamidases and glycosidases found on the API 32A test strip are also found on the less expensive AP1-ZYM strip which is designed to allow semi-quantitation of enzymatic activity. These tests are quite useful for directly differentiating a wide range of strains from each other, particularly if high levels of discrimination are required. All of the test strips can be easily set up using suspensions of cultures in sterile seawater and incubating the strips at about 10°C (or lower if necessary) for several days. Subsequent analysis and interpretation of the test strips should be performed according to the lnanufacturer's instructions.
Hydrolysis of complex and simple substrates Proteins
Gelatin hydrolysis. Gelatin hydrolysis can be tested in two ways.The first way is more subject to error especially if plates are incubated too long, however, the test is very simple and does not require any special preparation. Dissolve I% gelatin in basal growth media and pour as plates. Following incubation, plates are flooded with I M HCI to precipitate unhydrolysed gelatin that appears white, while clear zones around the growth are indicative of hydrolysis. It is critical that the plates are not incubated for too long (for most psychrophiles up to 5-7 d at I 0°C) as the gelatin hydrolysis zones will quickly cover the entire plate. An alternative method is to use commercially available sterile gelatincharcoal discs (Oxoid) which are added directly to the sterile liquid basal media. As gelatin hydrolysis occurs the charcoal is released into the media.
607
•
Casein hydrolysis. A 10-20% suspension of casein powder or skimmed milk in distilled water is autoclaved at reduced temperature (I 10°C, 20 min) and is then added to an equal volume of sterile basal agar medium. It is important not to autoclave the casein with the basal media as the casein interacts with the agar and precipitates. • Elastin and fibrinogen hydrolysis. Both proteins should be tested as thin overlays (0.5%-I% of protein in the basal agar medium) over a base made of unsupplemented agar. Hydrolysis of the proteins usually occurs within 7-14 days incubation indicated by the appearance of clear zones around growth.
Hydrolysis of polysaccharides The hydrolysis of polysaccharides such as starch, chitin, alginate and agar are abilities c o m m o n a m o n g marine bacteria and to various saccharolytic psychrophilic bacteria. •
Starch hydrolysis. Starch hydrolysis can be simply tested by supplementing the basal growth agar medium with I% starch and sterilizing the medium at reduced temperature (I 10°C, 20 min). Following sufficient incubation (at least 7 days) plates are flooded with a 1:5 dilution of Lugol's iodine solution (I g KI and I g sublimed iodine in 100 ml distilled water).The areas of the medium containing unhydrolysed starch are stained dark purple while hydrolysed zones around growth are clear. • Chitin hydrolysis. The test requires prior purification of commercial practical grade crab shell chitin; however, purified chitin can be purchased but it is prohibitively expensive. Add the precipitated purified chitin as a thin overlay in a mineral salts agar medium to achieve about a I% (w/v) concentration. Chitinase activity is indicated by clear zones around the growth. Chitin agar: 15 g agar, 3 g precipitated chitin, 2 g (NH4)2SO4, 0.7 g KH2PO4, I mg FeSQ and I mg MnSO, added to 1000 ml artificial seawater. • Alginate hydrolysis. Supplement the basal medium with I% sodium alginate (add with vigorous stirring and heating) and add about I% agar to form a solid medium. Hydrolysis is indicated by clearing zones around the growth. • Agar hydrolysis. Hydrolysis is indicated by softening, pitting or liquefaction of the agar medium surrounding and beneath growth.
Lu~ol's iodine solution. Grind 1 g of KI and i g of sublimed iodine in a mortar while adding small amounts of water. Once an even solution is formed dilute the iodine solution to 100 ml. Store in the dark.
Chitin purification. Add 40 g of chitin to 400 ml of cold concentrated HC1 and then precipitate the chitin by adding the solution to 2 1 of distilled water at about 5°C. Filter the suspension through W h a t m a n no. 1 filter paper. Re-suspend the chitin in distilled water and dialyse against tapwater overnight. Adjust the p H to 7.0 using KOH.
Lipolytic enzymes Esterase activity. Tweens 20, 40, 60, 80 (esters of myristic, stearic, palmitic and oleic acids, respectively) or tributyrin can be used as substrates for 608
esterases. For this Tweens are prepared as 10°~ solutions in distilled water and sterilized separately from the basal agar media preventing precipitation. Different Tweens are then added to the sterile molten agar to obtain a 1~/~ concentration and mixed to fully disperse the Tween. Tributyrin can be added directly to the medium (supplemented with 1 ~;{ polyvinyl alcohol to aid dispersal) before autoclaving, to achieve a final concentration of 1~/~. Hydrolysis of the Tweens is indicated by an opaque hazy zone of calcium soap crystals surrounding the growth. Hydrolysis of tributyrin is indicated by clear zones appearing in the initially cloudy medium.
Lecithinase activity. Lecithinase activity can be tested by adding to the basal medium, sterile egg yolk emulsion (about 5~/~final concentration) (Oxoid) and observing for opaque zones over and surrounding growth. Lipase activity. True lipase activity using olive oil or cottonseed oil (Sigma), inexpensive triglyceride substrates (other similar plant oils can be used), can be detected most directly using the procedure of Kouker and Jaeger (1987). To sterile molten basal agar medium, held at about 50°C, add a solution of 2.5% olive oil and 0.001(/~ rhodamine B (31.25 ml 1 ' of media) with vigorous shaking. After standing to allow the foaming to subside the medium is poured as plates. After at least 14 days incubation the plates are observed under long wavelength UV light (about 250 am) (e.g. using a hand-held or normal transilluminator or by using a UV light box). Strains producing a lipase develop a bright orange fluorescence. Strains not producing lipase produce no fluorescence. Other compounds Tyrosine hydrolysis or uricase activity can be tested by adding 1% tyrosine or 1'7~ uric acid to the basal agar media and observing for clearing zones around growth. Certain species on tyrosine agar will also produce red-brown diffusible pigments that are catabolites from the tyrosine degradation and can be used as an extra level of characterization. Activity for [3-glucosidase can be assayed by testing for the hydrolysis of esculin. To do this add 0.1% esculin and 0.01% ferric citrate to the basal agar medium. Esculin hydrolysis is indicated by the appearance of dark tan pigment diffusing into the agar. The production of deoxyribonuclease can be tested using DNAse Test agar (Oxoid),which is prepared in natural or artificial seawater and supplemented with 0.01 g toluidine blue. DNAse activity is indicated by the media around growth changing to red while negative strains stay blue.
Oxidation/fermentation of carbohydrates
The medium of choice for testing the ability of marine bacteria, including psychrophiles, to acidify carbohydrates is the Leifson O / F medium which consists of: 1 g casitone, 0.1 g yeast extract, 0.5 g ammonium sulfate, 0.5 g 609
tris buffer, 0.01 g phenol red and 3 g agar in 1000 ml natural or artificial seawater with p H adjusted to about 7.0. The ability to oxidize and ferment carbohydrates can be tested by first dispensing the basal Leifson m e d i u m into 20 ml screw cap tubes (two tubes per strain) and autoclaving. For sugars which are heat labile (e.g. most mono- and disaccharides) these should be a d d e d following autoclaving from filter-sterilized 20% stock solutions to achieve a concentration of 0.5-1%. The tubes are inoculated by stabbing them with an inoculating wire right to the base of the tube. One tube is then sealed with sterile liquid paraffin or with molten 3% agar. Acidification is indicated by the m e d i u m color changing from red to yellow. The presence of fermentation is indicated by the bottom of the tube or the entire tube turning yellow in the sealed tube. A strictly oxidative organism produces a distinct color at the top of the tube of the sealed tube. For testing only oxidative acid production from carbohydrates, agar plates (made with 1.5% agar) can be used instead of tubes.
Carbon source and nutritional tests
The utilization of sole carbon sources for carbon, energy and in some cases for nitrogen must be tested in a m e d i u m that is defined sufficiently for bacterial growth. Many marine bacteria can grow in either liquid or agar media containing seawater, a simple combined energy source and a carbon substrate while others require extensive supplementation including a d d e d vitamins, yeast extract, amino acids and possibly other growth factors. This may lead to problems if a large mix of strains is being investigated as some strains m a y be able to grow quite well on supplemented media, e.g. oligotrophic growth. Thus suitable controls lacking supplements and lacking carbon sources are very important. Obviously, strains able to hydrolyse agar must be tested in liquid media. A useful seawater mineral salts media broadly applicable to test psychrophiles consists of the following: 1 g a m m o n i u m chloride, 0.1 g yeast extract and 2 ml H u t n e r ' s mineral salts (see above) dissolved in 1000 ml of natural or artificial seawater. Carbon sources should then be a d d e d at a concentration of 0.1%, except carbohydrates which should be added at 0.2%. Labile and volatile substrates should be filter sterilized before addition to the sterile basal medium. Following addition of carbon sources the p H may need adjustment to about 7.0-7.5. After autoclaving, 10ml of vitamin solution stock is then a d d e d to media (cooled to about 50°C). If agar is being used for media, a high purity grade (Agarose, Agar Noble) should be used to reduce background growth. Incubation should proceed for up to 1 m o n t h at 10°C with close comparison m a d e with control plates lacking a carbon source.
References
Balch, W. E., Fox, G. E., Magrum, L. J., Woese, C. R. and Wolfe, R. S. (1979). Methanogens: reevaluation of a unique biological group. Microbiol. Rev. 43, 260-296.
610
Bligh, E. G. and Dyer, W. J. (1959). A rapid method of total lipid extraction and purification. Call. J. Biochem. Physiol. 37, 911-917. Bowman, J. P., McCammon, S. A., Brown, J. L., Nichols, P. D. and McMeekin, T. A. (1997a). Psychroserpens burtonensis gen. nov., sp. nov., and Gelidibacter algeiTs gen. nov., sp. nov., psychrophilic bacteria isolated from Antarctic lacustrine and sea ice habitats, h#. J. Syst. Bacteriol. 47, 670-677. Bowman, ]. P., McCamnron, S. A., Brown, M. V., Nichols, D. S. and McMeekin, T. A. (1997'o). Diversity and association of psychophilic bacteria in Antarctic Sea ice. Appl. EHviroH. Microbiol. 63, 3068-3078. Bowman, J. P., McCammon, S. A., Nichols, D. S., Skerratt, J. H., Rea, S. M., Nichols, P. D., and McMeekin, T. A. (1997c). Shewnuella gelidimariua sp. nov. and ShewalTella h'igidimarina sp. nov., novel Antarctic species with the ability to produce eicosapentaenoic acid (20:5o)3) and grow anaerobically by dissimilatory Fe(Ill) reduction, h#. J. Syst. Bacteriol. 47, 1040-1047. Bowman, J. P., McCammon, S. A. and Skerratt, J. H. (1997d). Methylosphaera hansonii gen. nov., sp. nov., a psychrophilic, group I methanotroph from Antarctic marine salinity; meromictic lakes. Microbiology 143, 1451-1459. Bowman, J. P., Gosink, J. J., McCanrmon, S. A., Lewis, T. L., Nichols, D. S., Nichols, P. D., Skerratt, J. H., Staley, J. T. and McMeekin, T. A. (1998a). Novel Colwellia species isolated from Antarctic fast ice: psychrophilic, marine bacteria with the ability to synthesize docosahexaenoic acid (22:60)3). hlt. 1. Syst. Bacteriol. 48, 1171--1180. Bowman, J. p., McCammon, S. A., Brown, J. L. and McMeekin, T. A. (q998b). Glaciecoh7 putIicen gen. nov., sp. nov. and Glaciecola pallidula gen. nov., sp. nov.: psychrophilic, marine bacteria from Antarctic sympagic habitats, hH. J. Syst. B~Tcteriol. 48, 1213-1222. Bowman, J. P., McCammon, S. A., Lewis, T. L. and Nichols, D. S. (1998c). Description of Psychroflexus torquis gen. nov. sp. nov., a psychrophilic bacterium from Antarctic Sea ice with the ability to form polyunsaturated fatty acids and the reclassification of Flavobacterium gondwalletlse Dobson, Franzmann 1993 as Psychroflextts ,\~oJidwalletlse gen. nov. comb. nov. Microbiology 144, 1601 1609. Bowman, J. P., Rea, S. M., Brown, M. V., McCammon, S. A. and McMeekin, T. A. (1998d). Investigation of aspects of community structure and psychrophily in Antarctic microbial ecosystems. Procecdiuss off the 8th IHterluHio~uTlSymposium oH Microbial Ecolo\,y, Halifax, Nova Scotia. Cohen-Bazire, G., Sistrom, W. R. and Stanier, R. Y. (1957). Kinetic studies of pigment swlthesis by nonsulfur purple bacteria. ]. Cell. Comp. Physiol. 49, 25-68. Dalgaard, P., Ross, T., Kamperman, L., Neumeyer K. and McMeekin, T. A. (1994). Estimation of bacterial growth rates from turbidimetric and viable count data. h#. J. Food Miclvbiol. 23, 391-404 Delille, D. (1996). Biodiversity and function of bacteria in the Southern Ocean. Biodivers. CoHserv. 11, 1505-1523. Deming, J. W., Somers, L. K., Straube, W. L., Swartz, D. G. and MacDonald, M. T. (1988). Isolation of an obligately barophilic bacterium and description of a new genus, Colwellia gen. nov. Syst. Appl. Microbiol. 10, 152-160. Dunkelblum, E., Tan, S. t t. and Silk, P. J. (1985). Double-bond location in monounsaturated fatty acids by dimethyI disulphide derivitization and mass spectrometry: application to analysis of fatty acids in pheromone glands of four lepidoptera. I. Chem. Ecol. 11, 265-277. Fay, L. and Richli, U. (1991). Location of double bonds in polyunsaturated fatty acids by gas chromatography-mass spectrometry after 4,4-dimethyloxazoline derivitization. ]. Chromatogr. 541, 89-98.
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Feller, G. and Gerday C. (1997). Psychrophilic enzymes - molecular basis of cold adaptation. Ceil. Mol. Life Sci. 53, 830 841. Feller, G., Narinx, E., Arpigny, J. L., Aittaleb, M., Baise, E., Genicot, S. and Gerday, C. (1996). Enzymes from psychrophilic organisms. FEMS Microbiol. Rev. 18, 189-202.
Franzmann, P. D. and Dobson, S. J. (1992) Cell wall-less, free-living spirochetes in Antartica. FEMS Microbiol. Lett. 97, 289-292. Franzmann, P. D, Liu, Y., Balkwill, D. L., Aldrich, H. C., Conway de Macario, E. and Boone, D. R. (1997). Methallogeuiun~ fri\qdum sp. nov., a psychrophilic, H-,using methanogen from Ace Lake, Antarctica. Int. J. Syst. Bacteriol. 47, 1068-1072. Gibson, A. M., Bratchell, N. and Roberts, T. A. (1987). The effect of sodium chloride and temperature on the rate and extent of growth of Clostridium botulinum type A in pasteurized pork slurry. J. Appl. Bacteriol. 62, 479-490. Gosink, J. J., Herwig, R. P. and Staley, J. T. (1997). Octadecabacter arcticus gen. nov., sp. nov., nonpigmented, psychrophilic gas vacuolate bacteria from polar sea ice and water. Syst. Appl. Microbiol. 20, 356-365. Gosink, J. J., Woese, C. R. and Staley, J. T. (1998). Polaribacter gen. nov., with three new species, P. irgensii sp. nov., P. fraHzmannii sp. nov. and f~ filamentus sp. nov., gas vacuolate polar marine bacteria of the Cytopha~a-Flavobacterium-Bacteroides group and reclassification of 'Fh'ctobacillus glomeratus' as Polaribacter glomeratus comb. nov. Int. J. Syst. Bacteriol. 48, 223 235. Helmke, E. and Weyland, H. (1995). Bacteria in sea ice and underlying water of the eastern Weddell Sea in midwinter. Mar. Ecol. Prog. Ser. 117, 269-287. Irgens, R. L., Suzuki, 1. and Staley, J. T. (1989). Gas vacuolate bacteria obtained from marine waters of Antarctica. Curr. Microbiol. 18, 261 265. Irgens, R. L, Gosink, J. J. and Staley, J. T. (1996). Polaromonas vacuolata gen. nov., sp. nov., a psychrophilic, marine, gas vacuolate bacterium from Antarctica. Int. /. Syst. Baeteriol. 46, 822-826. Knoblauch, C., Sahm, K. and Jorgensen, B. B. (1999). Psychrophilic sulfatereducing bacteria isolated from permanently cold Arctic marine sediments: description of Desulfofrigus oceane1~se gen. nov., sp. nov., Desulfofrigus fragile sp. nov., Desulfofaba go/ida gen. nov., sp. nov., Desulfotalea psychrophila gen. nov., sp. nov. and Desulfotalea arctica sp. nov. Int. J. Syst. Bacteriol. 49, 1631-1643. Koch, A. L. (1994). Growth measurement. In: Methods for General and Molecular Bacteriology (P. Gerhardt, R. G. E. Murray, W. A. Wood and N. R. Krieg, Eds), pp. 257-260. American Society for Microbiology, Washington, DC. Kouker, G. and Jaeger, K.-E. (1987). Specific and sensitive plate assay for bacterial lipases. Appl. Environ. MicJvbiol. 53, 211-213. Morita, R. Y. (1975). Psychrophilic bacteria. Bacteriol. Rev. 39, 144-167. Mountford, D. O., Rainey, E A., Burghart, J., Kaspar, H. E and Stackebrandt, E. (1998). Psychromonas antarcticus gen. nov., sp. nov., a new aerotolerant anaerobic, halophilic psychrophile isolated from pond sediment of the McMurdo Ice Shelf, Antarctica. Arch. Microbiol. 169, 231-238. Nichols, D., Bowman, J., Sanderson, K., Mancuso Nichols, C., Lewis, T., McMeekin, T. and Nichols, P. D. (1999). Developments with Antarctic microorganisms: culture collections, bioactivity screening, taxonomy, PUFA production and cold-adapted enzymes. Curt. Opiu. Bioteclmol. 10, 240-246. Nichols, D. S. and Russell, N. J. (1996). Fatty acid adaptation in an Antarctic bacterium - changes in primer utilization. Microbiology 142, 747-754. Nichols, D. S., Nichols, P. D. and McMeekin, T. A. (1995). Ecology and physiology of psychrophilic bacteria from Antarctic saline lakes and sea-ice. Sci. Proy,. 78, 3ll-347.
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Nogi, Y. and Kato, C. (1999). Taxonomic studies of extremely barophilic bacteria isolated from the Mariana Trench and description of Moritella yayaHosii sp. nov., a new barophilic bacterial isolate. Extremophiles 3, 71-77. Nogi, Y., Kato, C. and Horikoshi, K. (1998a). Moritella japonica sp. nov., a novel barophilic bacteriurn isolated from a Japan Trench sediment. J. Gen. Appl. Microbiol. 44, 289-295. Nogi, Y., Kato, C. and Horikoshi, K. (1998b). Taxonomic studies of deep-sea barophilic Shewanella strains and description of Shewanella violacea sp. nov. Arch. Microbiol. 170, 331 338. Ratkowsky, D. A., Lowry, R. K., McMeekin, T. A., Stokes, A. N. and Chandler, R. E. (1983). Model for bacterial culture growth rate throughout the entire biokinetic temperature range. J. Bacteriol. 154, 1222-1226. Russell, N. J. and Nichols, D. S. (1999). Polyunsaturated fatty acids in marine bacteria--a dogma rewritten. Microbiology 145, 767-779. Urakawa, H., Kita-Tsukamoto, K., Steven, S. E., Ohwada, K. and Cohvell, R. R. (1998). A proposal to transfer Vibrio marimts (Russell 1891) to a new genus Moritella gen. nov. as Moritella marilu7 comb. nov. FEMS Microbiol. Lett. 165, 373-378. White, D. C., Davis, W. M., Nickels, J. S., King, J. D. and Bobbie, R. J. (1979). Determination of the sedimentary microbial biomass bv extractible lipid phosphate. Oecologia 40, 51-62. ZoBell, C. E. (1946). Matqne Microbiology. Chronica Botanica Co., Waltham, MA, USA.
List of suppliers Alltech Associates, Inc. 2051 Waukegan Road Deerfield IL 60015, USA Tel: +1-847-948-8600 Fax: +1-847-948-1078 http://alltechweb.com
bioM6rieux, Inc. 595 Anglum Road Hazelwood MO 63042-2320, USA Teh +1-314-731-8500 Fax: +1-314-731-8700 http: //www.bionlerieux.con'l
G C vials, GC c o l u m n s
API test strips
Becton-Dickinson Biociences (Difco Products) 7 Loveton Circle Sparks MD 21152, USA Tch +1-410-416-3000 (central operator) +1-800-638-8663 (USA only) http://www.bdms.com/difco/index.html
Fluke Corporation 6920 Seaway Boulevard Everett WA 98203, USA Tel: +1-425-347-6100 Fax: +1-425-356-5116 http://www.fluke.coni
M a r i n e 2216 agar, microbiological m e d i a reagents (yeast extract, p e p t o n e , etc.), agar noble
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Electronic h a n d - h e l d t h e r m o m e t e r with thermocouple
Hewlett-Packard Corporation 3000 Hanover Street Palo Alto CA 94304-1185, USA Tel: +1-650-857-1501 Fax: +1-650-857-5518 h ttp://www.hp.com/
GC-MS, GC columns
Sigma-Aldrich Chemical Co. P.O. Box 14508 St. Louis MO 63178, USA Teh +1-314-771-5750 +1-800-521-8956 Fax: +1-314-771-5757 +1-800-325-5052 http://www.sigma-aldrich.cottl
Artificial sea salts; chemicals for reagents, buffers and media; filter paper
Mallinckrodt Inc. 675 McDonnell Boulevard Hazelwood MO 63042, USA Tel: +1-314-654-2000 Fax: +1-314-654-2328 http://www.mallinckrodt.com/
Nanograde organic solvents Oxoid, USA Inc. 800 Proctor Avenue Ogdensbu~ NY 13669, USA Tel: +1-800-567-8378 Fax: +1-613-226-3728 http://www.oxoid.com/
Spectronic Instruments Inc. 820 Linden Aveue Rochester NY 24625, USA Tel: +1-716-248-4000 +1-800-654-9955 Fax: +1-716-248-4014 http: //www.spectpvnic.com/
Spectronic spectrophotometers that can read test tubes
R2A agar, microbiological media reagents (yeast extract, peptone etc.), egg yolk emulsion, gelatincharcoal disks, triple-sugar iron agar, DNAse test agar
Toyo Roshi Kaisha Ltd., Advantec 1510, Bldg., 1-5-10 Kotobuki, Taito-ku Tokyo 111-0042, Japan Teh +81-3-3842-6290 Fax: +81-3-3842-6299 http://www.advm~tec.co.jp/
Temperature gradient incubator
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30 Deep-sea Piezophilic Bacteria A Aristides Yayanos University of California San Diego, Scripps Institution of Oceanography,La Jolla, CA 92093-0202, USA
CONTENTS Introduction Sources of information on high pressure technique Sampling the deep sea Pressure vessels,fittings,gauges, and pumps Culture containers Assay for colony-forming abilityat high pressure Enrichment cultures at high pressure Maintenance of piezophilic bacteria High pressure laboratory instruments for growth characterization Pressurized temperature gradient incubators Safety issues with high pressure equipment
~,4,~,~,~,~, I N T R O D U C T I O N "That, nevertheless, the microbiologist so ~]:ten succeeds in isolating spec!fic microbes from a given sample ~ff soil or water is due to a methodological principle first applied by Winogradsky, and still more consciously developed with quite amazing success by my great predecessor in the Delft chair of microbiology, Beijerinck. This principle has been dubbed by Beijerinck the ecological approach; its application depends on a well-considered selection of the conditions in a primary culture medium, thus causing ptvferential growth of a certain type of germ, ultimately leading to a predominance of the conditionally fittest. As soon as this stage is attained, isolation qf the prevalent organism with the aid of methods commonly used by mictvbiologists is, of course, an easy affair" (Kluyver and van Niel, 1956). The ecological approach (also k n o w n as the enrichment culture technique) praised so highly by Kluyver is part of the experimental backbone of microbiology. The decisive incorporation of pressure into the ecological approach begins with the work of ZoBell, Johnson, Morita and O p p e n h e i m e r (ZoBell, 1952; Yayanos, 2000). Pressure is seen widely today as a significant environmental parameter essential to research on inhabitants of the deep ocean, the seafloor beneath it, and continental subsurface environments. With reference to work in a high-pressure setting, we METHODS IN MICROBIOLOGY,VOLUME30 ISBN 0-12-521530-4
Copyright © 2001 Academic Press Ltd All rights of reproduction in any form reserved
should modify Kluyver's statement, to read that 'the aid of methods commonly used by microbiologists' is not always 'an easy affair.' Although efforts to simplify high-pressure microbiological methods are paying off, they remain relatively clumsy compared to methods used at atmospheric pressure alone. Through research over the past 25 years, we have a good idea of circumstances where high-pressure methods must be used and of those where high-pressure methods either can be set aside confidently or used in control experiments as required. In this context, a summary of the present status is: 1. Hyperpiezopsychrophiles (Yayanos, 1995) are psychrophilic microorganisms having a maximum growth rate at a pressure greater than 50 MPa and growing poorly, if at all, at atmospheric pressure. They are the most pressure adapted bacteria known. Many of these pressureadapted bacteria survive brief decompressions. Thus, their cultivation is possible starting with unwarmed, decompressed samples even of the greatest ocean depths (beyond 10 500 m) if the enrichment cultures are compressed within an hour or so of the sample's arrival on board ship. Hyperpiezopsychrophiles from a 6000 m depth survive in water samples kept on ice at atmospheric pressure for two weeks (Yayanos and DeLong, 1987). 2. High-pressure laboratory methods are indispensable for pure-culture deep-sea microbiology. Enrichment cultures incubated at the pressure and temperature of that depth are likely to yield true inhabitants of that depth. The conduct of enrichments at high pressure not only facilitates and enables the growth of authochthonous microorganisms, but also inhibits the growth of allochthonous ones. The warmer a deep-sea habitat is, the less decompression sensitive its inhabitants are (Yayanos, 1995). Nevertheless, enrichment cultures at high pressure can yield new organisms from these warm habitats as clearly shown in a recent study on a hydrothermal vent sample (Marteinsson et al., 1999). 3. Sampling the deep ocean with pressure-retaining devices is essential to the search for deep-sea microorganisms that die instantly upon decompression. Although such organisms have not yet been found, they probably exist. 4. Future research with pressure-retaining samplers will give us a better understanding of how pressure change affects the activity of communities of microorganisms. The type of pressure change in need of further study is not only that imposed on a community when we remove it from the ocean, but also that caused by natural processes. For example, particle sedimentation and animal vertical migration cause attached microbial communities to experience pressure change. Furthermore, natural communities contain mixtures of allochthonous and autochthonous cells that will respond differently to imposed pressure changes. I do not describe herein methods for the deep ocean collection of material in pressure-retaining devices. At this time, these methods require a 616
substantial financial investment in equipment and are prototypical rather than conventional methods. The laboratory methods described in this chapter are useful for the enrichment, isolation, and pure culture study of some of the microbial inhabitants of deep-sea and subsurface habitats. The methods are accessible to anyone wishing to make a modest investment in high-pressure equipment.
4, 4, 4, 4, 4, 4, S O U R C E S O F I N F O R M A T I O N PRESSURE TECHNIQUE
ON HIGH
The background for working with high-pressure technique in the laboratory derives largely from the field of chemical engineering where industrial needs fostered progress in the 20th century (Newitt, 1940; Dodge, 1950; Comings, 1956; Tongue, 1959). Other reviews include: Johnson et al. (1954), Tsiklis (1968), Suzuki (1973), Hawley (1978), and Holzapfel and Isaacs (1997). Fryer and Harvey (1998) is an excellent resource for pressure vessel design. Several of these books and articles are also sources of information on safety issues in high-pressure technique. Four recent reviews contain material relevant to high pressure microbiological methods (Ludlow and Clark, 1991; Deming, 1997; Yayanos, 1998; Horikoshi and Tsujii, 1999).
SAMPLING
THE DEEP SEA
The discovery and cultivation of new microbial species and the study of microbial activities in natural settings are the two principal reasons for the collection of samples from the deep sea. There are devices that retrieve water (Macdonald and Gilchrist, 1969; Jannasch et al., 1973; 1982; Tabor et al., 1981; Bianchi et al., 1999), sediments (Murray et al., 1989), and animals (Macdonald, 1978; Yayanos, 1978) while keeping them at high pressure. An absolute requirement is to avoid the warming of any sample collected in the cold deep ocean as you bring it to the ship and process it in the laboratory. Another recommended practice is to prevent the exposure of samples and cultures to UV light (from sunlight or from laboratory lighting). Deep-sea bacteria examined to date show extreme sensitivity to UV exposure. Whereas pressure-retaining deep-ocean samplers are available only as prototype instruments, other samplers (Niskin bottles, Go-Flo bottles, box corers, gravity corers, and sediment traps) are more widely available. The recurring question in all microbiological sampling is whether decompression of the sample is acceptable. There is no generic answer. Rather, the outcome of a given deep-sea sampling program needs to be evaluated for a possible influence of decompression. If the purpose of a sample is to determine the activity of a natural microbial community, then pressure617
retaining samplers are essential. If the purpose of deep-sea sampling is to obtain some interesting representatives of the true inhabitants of the deep sea, then decompressed samples kept at deep-sea temperatures during retrieval provide viable cells for laboratory enrichments. These incubations, however, must be done at the habitat pressure and temperature as well as at appropriate nutrient levels and redox conditions. The methods in the remainder of this chapter are for the laboratory study of deep-sea microorganisms.
eee,ee
PRESSURE VESSELS, FITTINGS, GAUGES,AND PUMPS Several vendors provide a wide selection of pressure vessels in full compliance with safety codes. The reasons for undertaking an in-house fabrication of vessels include the need for a custom design or for economy. Cost reduction is possible when fabricating a large number of pressure vessels. The book by Fryer and Harvey (1998) is an excellent guide for designing pressure vessels. If you choose to fabricate your own pressure vessel, determine if the materials you select are available in a pressure vessel grade. For use in marine microbiology, pressure vessel materials must withstand exposure to seawater and have neither an effect on the microorganisms in the vessel nor be affected by them. Type 316 stainless steel and titanium, particularly the alloy 6AI4V (6% aluminum and 4% vanadium), exhibit excellent corrosion resistance in seawater. Additional considerations arise because of the type of microorganism under study. For example, hydrogen-utilizing bacteria present a special problem because hydrogen interacts with many pressure vessel materials to weaken them (Vodar and Saurel, 1963; Ludlow and Clark, 1991). Experiments on deep-sea bacteria are seldom conducted at pressures beyond 100 MPa. This is important to keep in mind because the higher the working pressure the more expensive the pressure vessel. An additional benefit of working at pressures <110 MPa is that ~-inch tubing (0.030 inch i.d.) and fittings are entirely adequate. The flexibility o f " inch tubing enables connections between the various components of a high-pressure apparatus that are easily made in minutes. In comparison, setting up an apparatus with ~-inch tubing can require several days of cutting, threading, and bending tubes to join various components of a pressure apparatus.
Pressure vessels useful for microbiology Two types of pressure vessel are shown in Figures 30.1 and 30.2 in sufficient detail to allow their fabrication. The pressure vessel shown in Figure 30.1a and 30.1b is useful for incubating enrichment cultures and pour tubes, and for studying microbial growth kinetics. The design goals for
618
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Figure 30.1a. Drawing of pin-closure pressure vessel in sufficient detail to allow its fabrication (Yayanos, 1982).
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Figure 30.1b. Photograph of a pin-closure pressure vessel. Below it is a tubulated plastic culture bag containing glass balls.
the pressure vessel s h o w n in Figure 30.1 are as follows: to a c c o m m o d a t e a culture v o l u m e of at least 200 cm~; to allow for a relatively inexpensive fabrication; to be compatible with seawater; and, to p r o v i d e for quick access to its contents. The pressure vessel in Figure 30.1 is a piston-seal pin-closure vessel. The first design of a pressure vessel with a pin closure
619
Ti-GAL4V FBR AND
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Figure 30.2. Drawing of a pressure vessel ideal for small incubations and transport of field samples. The weakest part of the vessel is the wall at the thread relief. A slightly thicker wall is recommended for routine use of this vessel to 100 MPa.
w a s b y Gasche (Gasche, 1966) of Autoclave Engineers, Inc. (Erie, PA). Pinclosure vessels can be p u r c h a s e d from t h e m today. There are several differences b e t w e e n the vessel s h o w n in Figure 30.1 (designed w i t h o u t examining Gasche's design) a n d that of Gasche. One is that the closure seal for the vessel in Figure 30.1 is s i m p l y an O-ring piston seal. The vessel of Gasche has a metal to metal seal backed b y an O-ring that allows m u c h higher w o r k i n g pressures than a piston seal does. Another difference is that the pin used in Gasche's design has a handle on it. A third difference is that the top of the closure to the vessel of Gasche rests flush with the top of the vessel cylinder. The closure has a shoulder that sits on a ledge internal to the pressure vessel. In contrast, the pressure vessel in Figure 30.1 has a shoulder that rests on top of the vessel. The a d v a n t a g e s of this external shoulder are that it can be g r a s p e d to r e m o v e it and that m o r e pressure vessel material is available to s u p p o r t the pin. A d i s a d v a n t a g e is that the piece of bar stock from which the closure is m a d e is larger than that for a Gasche vessel of the s a m e size. In the use of any pin-closure vessel, the pin m u s t be inserted with care so that the ends of the pin are
620
flush with the sides of the vessel. Once a vessel is u n d e r pressure, the pin remains firmly in place. These vessels should never be pressurized without a pin in place because the cap could b e c o m e a projectile. A large r u b b e r b a n d a r o u n d the b o d y of the pressure vessel can be slipped over the pin to insure that it does not fall out w h e n the vessel is at a t m o s p h e r i c pressure. An i m p o r t a n t detail for the fabrication of a pin-closure pressure vessel is to insure that the pin material has a high shear strength. The pin of the vessel s h o w n in Figure 30.1 is of type 17-PH stainless steel that acquires high shear strength w h e n heat-treated. This pressure vessel can be safely used to 100 MPa. Significant cost savings accrue in each step of the fabrication of a vessel such as that s h o w n in Figure 30.1 w h e n a p p r o x i m a t e l y 50 or m o r e are fabricated concurrently. The steps for fabricating such a vessel are as follows: 1. Use bar stock of 2.5" d i a m e t e r m a d e of type 316 stainless steel (ASTM 479) for the cap and b o d y of the pressure vessel. Use bar stock of 0.75" d i a m e t e r m a d e of type 17-4 P H stainless steel for the pins. 2. Cut and face off bar stock for the b o d y of the vessels to a final length of 15" (or a n y desired length). 3. Gundrill bars to a d e p t h of 14" with a spherical radius at the b o t t o m and with a bore d i a m e t e r of 1.490" _+ 0.002". You will need to find a gundrilling shop and obtain a quotation for the cost of this step. 4. To a d e p t h of 2.75", hone (32 hone) the bore to an i.d. of 1.500" + 0.001". 5. To a d e p t h of l v.... , r e a m the bore to a d i a m e t e r of 1.530" and terminate the reamed portion with a 30 ° angle with the wall of the cylinder. 6. Drill v d i a m e t e r holes centered at 1~" from the bored end and p e r p e n dicular to the axis of the cylinder. 7. M a k e the cap per d i m e n s i o n s in Figure 30.1. The d i m e n s i o n s for the [ I1 F250C v high-pressure connection are in the catalogs of several m a n u f a c t u r e r s of high-pressure fittings. 8. M a k e the pins from the 17-4 PH bar stock. 9. Provide an O-ring of size 2-218. 10. Connect a t w o - w a y valve to the F250C connection in the cap with a nipple of vI '1 tubing. N o t e that the two seals in the valve are different. One is a metal-to-metal seal formed b y the tip of the valve stem and a seat in the valve body. The other seal is d u e to the packing a r o u n d the valve stem. M a k e sure that the metal-to-metal seal is closest to the pressure vessel. 11. Place a quick-connect nipple (described in the following section) on the other F250C connection on the valve. Pin-closure vessels o p e n easily and quickly. We connect the valve on a pressure vessel that is u n d e r pressure to a p u m p e q u i p p e d with a pressure gauge. Before o p e n i n g the pressure vessel, we raise the pressure to the level we expect to find. Then we slowly o p e n the valve on the pressure vessel. Usuall> the pressure on the g a u g e is within a few a t m o s p h e r e s of the expected value. We d e c o m p r e s s the pressure vessel b y o p e n i n g the relief valve on the p u m p i n g system. The d e c o m p r e s s i o n releases the load on the pin that we then easily p u s h out. Then, we pull off the lid of the
621
vessel by using a gentle back and forth twisting motion. Thus, we can open a pin-closure pressure vessel in five or ten seconds following its decompression. When the pressure in a vessel is either more or less than the expected value, the actual pressure can be estimated from a calibration graph. Of course, this procedure must be done for each pressure vessel size, p u m p i n g system, temperature, and hydraulic fluid. First, increase isothermally the pressure in a vessel to 30 MPa, for example, and close the pressure vessel valve. Second, increase the pressure in the p u m p i n g system to a target pressure of 50 MPa. The target pressure is the one we hypothetically expect to find in the pressure vessel. Third, slowly open the valve connecting the pressure vessel to the p u m p i n g system. The pressure on the gauge then drops to a value less than 50MPa and greater than 30 MPa. Repeat the above procedure for different pressures at a constant target value of 50 MPa. A graph of the gauge pressure versus the pre-set pressure in the vessel allows estimation of the actual vessel pressure if it drifted from the target value (50 MPa in this example) during the course of an incubation. The pressure vessel shown in Figure 30.2 is particularly suitable for fieldwork and for experiments involving small v o l u m e incubations. The closure shown in Figure 30.2 is one piece that has an O-ring groove and threads. One-piece closures, although not r e c o m m e n d e d for large pressure vessels, work very well with small ones. The pressure vessel shown in Figure 30.2 is small, light, corrosion resistant, easy to open, and easy to close. We decompress a small pressure vessel in the same way as described above for a pin-closure vessel. At atmospheric pressure, the threads no longer bear a stress and allow easy separation of the cap from the b o d y of the vessel. Since the vessel is small, turning it while holding the closure allows it to remain connected to the p u m p i n g system. Because the closure on this small vessel is a piston seal closure, it is unnecessary to tighten the closure to get a high-pressure seal. A highpressure seal will form as long as the cap screws onto the b o d y of the vessel to the m a x i m u m extent. Of course, the valve on the pressure vessel must be open to allow excess liquid in the pressure vessel to be expelled as you screw the closure onto the cylinder. Small vessels such as these are of great advantage in fieldwork because approximately 25 of them fit in an ice-filled small container to a final weight of less than 70 pounds. We have shipped such containers as excess baggage. Since these vessels are water filled and of a small v o l u m e (30 cm ~) they pose no hazard whatsoever to an aircraft or to baggage handlers. This statement does not apply, however, to water-filled vessels that contain liquid mercury, carbon monoxide, or other chemicals that m a y form reactive mixtures. Pressure vessels being shipped from the field can suffer decompression if the valve handles are b u m p e d . This type of decompression can be minimized by placing high-pressure plugs on each unoccupied valve connection. The female connection in the cap of the small field vessel is not shown in Figure 30.2. We usually use a 7~" high-pressure fitting. Valves for ~-" tubing are smaller than Ti /t valves and allow for a more robust connection than d o vI t, valves. 622
Quick-connect fitting Pin-closure vessels bestow a convenience on microbiology because they are easy to open and close without the use of tools. You can facilitate the process even more by eliminating the need for a wrench to connect and disconnect a pressure vessel to a p u m p i n g system. The quick-connect fitting (Yayanos and Van Boxtel, 1982) shown in Figure 30.3 consists of a male part (one for each pressure vessel) and a female part (one for each p u m p i n g system). The female part connects to a high-pressure p u m p i n g system via ~ inch tubing. The male part is a short nipple of " inch o.d. tubing threaded on one end and modified in two ways. First, seal the end of the nipple opposite the threaded end with a short stainless steel rod held in place by crimping the end of the tube, and silver solder. Next, drill a small hole through the nipple perpendicular to the cylindrical axis. The O-rings in the female part of the quick-connect fitting straddle this hole w h e n the female part is slipped over the nipple. Thereby, hydraulic fluid passes into or out of the pressure vessel. Each pin closure vessel has a modified nipple thereby facilitating its compression and decompression. In summary, the quick connect fitting allows mating of pressure vessels to a p u m p i n g system without the use of a tool. The pinclosure vessel along with the quick-connect fitting enables a variety of high-pressure microbiological experiments without the use of tools. The time saved is considerable w h e n accessing a large n u m b e r of pressure vessels in a day.
Figure 30.3. Photograph of a quick-connect fitting for ~-" high-pressure connections. A drawing to allow fabrication of this device can be found in Yayanos and Van Boxtel (1982).
623
Pressure gauges The measurement of pressure in experiments has errors that range from 1% to 0.01% of the full-scale pressure reading of the gauge. For microbiological work such as the isolation of new organisms or physiological studies, pressure gauges having a m a x i m u m pressure reading of 138 MPa and a pressure measurement error of _+1.38 MPa are entirely adequate. For studies of e n z y m e kinetics at high pressure, an analysis of systematic error will establish whether a gauge giving the highest possible accuracy in the pressure determination is necessary. Paro Scientific, Inc. manufactures a pressure gauge that allows determinations with an accuracy of •+0.01%. This accuracy is about as good as is possible to achieve. Finally, before placing an order for a pressure gauge, you need to determine whether the pressure gauge materials are compatible with the hydraulic fluid in y o u r experiments.
High-pressure pumps There are three types of high-pressure p u m p c o m m o n l y used. One generates pressure by driving a piston into a cylinder with a screw. Another type drives a piston into a cylinder by means of a hand-operated lever. A third type uses the action of air pressure on a large diameter piston to drive a small diameter piston into a liquid-filled cylinder. Screw-driven p u m p s are available from Ruska Instrument C o m p a n y and from High Pressure Equipment inc. Autoclave Engineers offers a well-made leveroperated pump. Haskel International, Inc. (Burbank, CA) manufactures a line of air-driven high-pressure pumps. A significant gain in convenience results from mounting a p u m p , gauge, and hydraulic reservoir beneath the top shelf of a laboratory cart. A long and flexible ~I outer diameter high-pressure line serves to connect the cart-mounted p u m p i n g system to pressure vessels on a laboratory bench or in an ice chest on the laboratory cart itself. PP
CULTURE
CONTAINERS
The selection or design of containers to confine cultures for incubation in pressure vessels is often a difficult problem in high-pressure microbiology. A culture incubated in a pressure vessel must remain axenic, be separate from the hydraulic fluid, and have the same pressure as the surrounding hydraulic fluid. Other desirable features for a culture container m a y include walls with a desired gas permeability, ports for sample removal, and space for stirring bars or marbles. The increasing variety of heat-sealable materials m a d e of flexible plastics is a technological advance of greatest significance for high-pressure microbiology because it makes possible the fabrication of culture containers with m a n y of the desirable features. The creative use of heat-sealing technology will continue to be of benefit to high-pressure microbiology. 624
Test tubes ZoBell a n d his colleagues, as others before them, confined cultures in glass test tubes sealed with rubber stoppers that transmitted pressure to the culture and p r e s u m a b l y g a v e satisfactory results. An easier w a y to seal test tube cultures is with Parafilm" slightly stretched and tightly w r a p p e d a r o u n d the m o u t h of the test tube, as previously described (Yayanos, 1980; Dietz and Yayanos, 1978).
Heat-sealable plastic transfer pipettes Polyethylene (PE) transfer pipettes come in a variety of sizes and in sterile packaging. Cultures are d r a w n into the bulb of a transfer pipette and m a n i p u l a t e d to r e m o v e air or other gas. The stem of the bulb is heat sealed with a n y of a variety of Teflon":-coated heat sealers. Sealed PE transfer pipettes are useful for confining s a m p l e s collected in the field, enrichment cultures, cultures with radiolabeled substrate, and inoculated solid media for the d e v e l o p m e n t of colonies. The long stem on the PE bulbs allows for collection of s a m p l e s over time. We r e m o v e a PE bulb from a pressure vessel, s w a b the stem with an ethanol w a t e r mixture (80~ v / v ) , and cut it just b e l o w the heat seal. Following the r e m o v a l of a desired a m o u n t of culture, w e again heat-seal the stem, p u t the bulb back into a pressure vessel, c o m p r e s s it, and r e s u m e the incubation. Figure 30.4 s h o w s a d r a w i n g of a heat-sealed bulb.
PE TRANSFER PIPET J J f.
-
, o
[
/ ,
/
COLONIES OF BACTERIA
HEAT SEAL
Figure 30.4. Drawing of a polyethylene (PE) transfer pipette or bulb filled with inoculated solid medium such as agar, silica gel, or gelatin and in which immobilized cells grew to form visible colonies. PE bulbs are excellent for pour tube assays and for obtaining a pure culture. A sterile syringe and needle are used to pierce the side of a PE bulb (after wiping the surface with ethanol) to pick a colony. PE bulbs allow for stab cultures. PE pipettes also serve in a similar way for the incubation of liquid cultures. Since PE is permeable to oxygen, aerobes will grow in these bulbs by surrounding them with oxygenated water. Strict anaerobes grow in PE bulbs by placing them in an oxygen impermeable bag filled with dithionite solution.
Plastic bags The first application of heat-sealed bags in microbiology was that of Berger and Tam (Berger and Tam, 1970). They grew Micrococcus euryhalis, a strict
625
aerobe, in a pressure vessel by placing the inoculated medium in a bag fabricated from oxygen-permeable silicone membranes. Oxygen enters such cultures from the surrounding hydraulic fluid. At that time, bags of silicone materials were fragile and did not become widely used. Subsequently, Helmke (Helmke, 1979) used gas-permeable polypropylene bags that are durable to confine cultures and to provide them with oxygen from oxygenated fluorocarbon hydraulic fluid in her studies of actinomycetes at high pressure. Teflon:' bags with high gas permeability are now also commercially available. Only through additional research will we be able to control the gas composition in high-pressure cultures for the full spectrum of biologically reactive gases. The following list is one of the procedures we use to fabricate plastic bags for high-pressure microbiology. 1. Purchase Kapak pouches (Stock no. 405) made of Scotchpak heatsealable polyester film (Kapak Corp., 5305 Parkdale Drive, Minneapolis MN 55414). These pouches are 10" by 12" and made of a 2 mil thick bilaminate of polyester and medium density polyethylene film. Kapak pouches made of 4 mil thick film are also available. While the oxygen permeability goes down as the thickness goes up, so does bag rigidity making them more difficult to work with. 2. Rinse the inside of a pouch with 95% ethanol followed by two rinses of deionized water (filtered through 0.2 lJm pore-size filters) and allow it to drain and dry in a laminar flow hood. This step removes residues of starch powder placed in the pouches by the manufacturer to reduce inner surface adhesion. Parenthetically, these bags are obviously not suited for studies of oligotrophic bacteria. 3. Heat-seal each pouch lengthwise at 2" intervals with a pneumatic impulse heat sealer (Stock no. 30-233, Accu-Seal Corp., 11055 Flintkote Avenue, San Diego, CA 92107). Cut tlle pouch along the middle of the "" heat-sealed beads to yield five culture bags per Kapak pouch. 4. Attach a polyethylene tube to one end of each pouch as follows. Place a 10" length of unenameled 24 gauge copper wire (Belden Wire and Cable, Richmond, IN 47375) through the bore of a 6" length of polyethylene tubing, 0.045" i.d. and 0.062" o.d. (lntramedic Stock no. 7431, Clay Adams/Becton Dickinson, Parsippany NJ 07054). Insert the tubing, with the wire extending about ~-" beyond its end, into an open end of the bag to a depth of 1". Seal the tubing into the bag with application of heat to both sides. At least ~" of the tubing should extend into the lumen of the bag. Apply sufficient heat to fuse the polyethylene tubing onto the inner surfaces of the bag so that there are no air bubbles or channels through the T bead. Check the seal with a dissecting microscope. After tile seal cools, withdraw the copper wire. Figure 30.5 shows the tubulated end of a bag. Note that tile formation of a leakproof seal between the polyethylene tubing and the bag becomes more difficult with wire sizes larger than 24 gauge. 5. Heat-seal the end of the polyethylene tubing. 6. Place about six glass or Teflon' balls of r diameter (Small Parts Inc., Miami, FL 33238-1966) into the bag if the experiment requires mixing the culture. A bag with glass balls is shown in Figure 30.1b. I ,1
I t,
626
Figure 30.5. Photograph of the tubulated end of a heat-sealed plastic culture bag.
7. Expose the bags for 30 min to UV light from a germicidal 15 W lamp with the bags placed 19 inches from the lamp. Whereas the polyester material is relatively opaque to 254 nm UV, the length of the exposure has been found to result in no bacterial contamination in experiments with deep-sea bacteria. We prepare for an experimental incubation by cutting open the end of the bag opposite the tubulation and pour or pipette a culture into the bag while holding or supporting it vertically in an ice bath. Alternatively, we place culture medium into a bag first and then add inoculum later. We periodically sample cultures via the tubing by first wiping it with 80-95% ethanol, allowing time for the residual ethanol to evaporate, and then cutting the end of the tubing with a flame sterilized scissors. We squeeze out a desired volume of culture, heat-seal the tubing, and resume the incubation.
Syringes Varieties of syringes have been used to isolate cultures from the hydraulic fluid in pressure vessels or from seawater in the deep sea. Confinement of cultures in syringes may be satisfactory for incubations lasting a few hours or days. Nevertheless, a potential problem with syringes is that fluid and cells can pass across the plunger seal. This is the main difficulty with syringes as culture containers. The extent of this concern is inadequately explored and largely ignored in many reports of high-pressure experiments based on syringe incubations. Where the use of a syringe is necessary or convenient, the choice is between purchasing or fabricating it. Purchased syringes usually require modification because they were not designed for use in pressure vessels. Many types of glass syringe usually
627
have a hollow plunger. A hole must be cut into the plunger to allow hydraulic fluid to enter it. Otherwise, the plunger will implode when it is compressed. The plunger on certain all-plastic syringes, such as those m a d e by Becton-Dickinson, consists of two pieces, one a plastic plunger frame and the other a rubber tip. The molded rubber tip has two circular ridges (providing O-ring like seals) between which there is a gap of air. The inside of the rubber tip rests on the plastic plunger frame in such a way that another pocket of air can become trapped between the inside of the rubber tip and the supporting plastic frame. When such syringes are used at atmospheric pressure, these air pockets cause the syringe plunger to seal more effectively than it would without them. For use in pressure vessels, however, it is important that these air pockets fill with hydraulic fluid. Otherwise the rubber tip will collapse during compression and the syringe will leak a portion of its contents into the surrounding hydraulic fluid and vice versa. Experiments show that leaks of this kind happen and that appropriately modified syringes do not leak their contents within the limit of detection by absorption spectrophotometry. Plastic syringes made by Sarstedt seem to work fine for microbiological studies, with no modification. Again, however, long duration experiments m a y give results of questionable value. A needle placed snugly over the tip of a syringe and poked into a silicone rubber stopper suffices to seal the tip. In summary, syringes that are designed for biomedical applications cannot be recommended as containers in high-pressure microbiological experiments if heat-sealable culture containers can readily be used instead. In addition to the above commercially available and appropriately modified syringes, we use the custom-designed syringe shown in Figure
TITANIUM CULTURE SYRINGE FOR HIGH TEMPERATURE/HIGH PRESSURE INCUBATIONS
Ti5OATitanium
tubing honed inside
0
~
TT
&5" .035"
2-213Viton o-ring w/backup(Teflon split)
.125"
Se,.....
1
1
Be.I
"
15-21AFINMA (HIP)
adapter
0.200" ~'q b~
1 '±
2OGV,IO °
1.15"
0.957'~..,IIi ~I J~ ~
~{ ~.200" }.q-.784"-~q
"Bleed 1/4-20eThread hl°
Beve~edgef " - ~' '~ 1.175" 1 178" 1175" 6AL4VTITANIUM PISTONANDENDPLUG
Figure 30.6. Drawing of a titanium syringe that is suitable for short duration incubations of cultures in pressure vessels. Any titanium alloy compatible with the microorganisms being cultured is suitable. Honing the inside of the tube is recommended. Lexan syringes are made from essentially the same drawing, but a thicker walled material is used for the syringe barrel. 628
30.6. The syringe has titanium parts allowing its use to high temperatures well above 373 K. The upper working temperature of this titanium syringe is a function of the O-ring materials used. This syringe was used in a study of Bacillus stearothermophilus and a study on the thermal decomposition of amino acids at temperatures above 523 K. For work on mesophilic and psychrophilic bacteria, we use a syringe of similar design fabricated from the autoclavable plastic LexaffL Custom-designed syringes have seals both at the tip and at the plunger that are more appropriate for high pressure use than those found on most commercially available syringes. Details of how to transfer culture between pressure vessels can be found in Yayanos et aI. (1983).
D i r e c t incubation in a pressure vessel Cultures can be put directly into a pressure vessel if it is made of a microbiologically compatible metal. There are few reports of studies using this technique. The most notable are the studies of natural populations of deep-sea bacteria that are incubated directly in pressure-retaining water samplers (Tabor et al., 1981; Jannasch et al., 1976). The samplers, however, have interior surfaces coated with Teflon''.
ASSAY FOR C O L O N Y - F O R M I N G A B I L I T Y AT H I G H PRESSURE Piezopsychrophilic bacteria from depths of approximately 3000m and greater grow poorly if at all when plated on nutrient agar plates incubated at atmospheric pressure. Two methods allow for colony growth in pressure vessels. One is the pour tube method and the other is the Doryaki method. Both methods have been used to isolate bacterial clones. The pour tube method serves very well to assay for colony-forming ability in experiments determining the sensitivity of cells to UV light, ionizing radiation, and chemicals (Yayanos, 1995).
Pour t u b e m e t h o d The pour tube method entails dispersing cells in a liquid just before it forms a gel. The cells become immobilized in the matrix of the gel. Nutrients diffuse to the immobilized cells that can grow and divide to form a visible colony of cells. Four types of solid media have been used to cultivate bacteria in pressure vessels. These are silica gel (Dietz and Yayanos, 1978), agar (Jannasch and Wirsen, 1984), gelatin (Yayanos and Dietz, 1983), and Gelrite (Deming and Baross, 1986). The first three of these solid media work well to isolate deep-sea psychrophilic bacteria. Gelrite was used to isolate thermophilic bacteria inhabiting hydrothermal vents. We find that gelatin gels are the easiest to use with cells that do not digest gelatin.
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Procedure for preparation of silicic acid sol, modified from that previously described by Dietz and Yayanos (1978) 1. Place 454 g of Amberlite ~ 120-H-C-P in a glass c o l u m n (4.4 cm d i a m eter, 43 cm long). 2. Pass 1 1 of N a n o p u r e ® w a t e r through the c o l u m n being careful to p r e v e n t the formation of discontinuities and the t r a p p i n g of air bubbles in the packed resin. If g a p s are present, back-flush the column with N a n o p u r e ®w a t e r and stir with a glass rod to r e m o v e a n y air. 3. Pass 250 ml of 4 N HC1 through the c o l u m n and as in all steps, use a flow rate of 100 ml m i n ' or less. The v o l u m e of resin will contract slightly as the regeneration proceeds. 4. Flush column with 2 1 of N a n o p u r e ®water to r e m o v e residual HC1. 5. A d d 500 ml 0.5 M NaSiOB.9H20 (71.05 g per 500 ml) to the c o l u m n a n d m a k e sure that solution level does not fall b e l o w the surface of the resin. 6. Feed N a n o p u r e ® w a t e r onto the c o l u m n and discard the first 200 to 250 ml of effluent. 7. Continue to feed N a n o p u r e ® w a t e r onto the column and collect the next 1000 ml of effluent into a polyethylene g r a d u a t e cylinder. 8. Transfer the effluent to a polyethylene beaker. 9. Adjust the p H of the solution to 1.5 b y a d d i n g HC1 (approximately 14 ml of 4 N HC1, d e p e n d i n g on the final concentration of the silicate ion). Use of m o r e concentrated HC1 m a y be desirable to m i n i m i z e the change in silicic acid concentration on addition of HCI. 10. Dispense into plastic screw cap E r l e n m e y e r flasks (500ml or 11 capacity) to no m o r e than two-thirds full. Flasks m a d e of p o l y p r o p y lene, p o l y c a r b o n a t e or p o l y m e t h y l p e n t e n e w i t h s t a n d autoclaving. Avoid the use of glass flasks because of the possibility that the s o d i u m or other metal ion residues on the glass m a y shorten the shelf life of the sol. 11. Autoclave the silicic acid sol at 15 psi for 20 min. Store until required at 2 to 4°C. 12. Wash the column with 11 of N a n o p u r e ® w a t e r i m m e d i a t e l y after use (step 7), and b a c k w a s h if necessary to r e m o v e t r a p p e d air. If gelation has occurred in the column, flush with 11 of 5% K O H followed b y a w a t e r rinse. 13. Regenerate the c o l u m n by flushing it with 500 ml of 4 N HC1 and store the c o l u m n in this solution.
Procedure for formation of inoculated silica gels 1. A d d the following solutions, all sterile and at 2 to 4°C, to 40 ml of silicic acid sol in a sterile 100 ml Tripour ®beaker on ice, and a d d with agitation or stirring: 2.3.0 ml 1 N N a O H 3.4.0 ml 10× TGYE (tryptone-glucose-yeast extract) m e d i u m 4.4.0 ml 10x ASW-MOPS (artificial seawater with buffer at desired pH) 5. The order of addition of the solutions to the silicic acid sol is critical in
630
order to avoid the formation of poor gels. Pre-measuring the NaOH, TGYE and ASW in snap-cap plastic tubes facilitates speed and ease of making the sol gels. 6. Immediately after Step 1 add, with agitation or stirring, up to 1 ml of cell suspension. 7. The mixture thickens in approximately 15 min and gels in 20 min. This gives ample time to stir the mixture and dispense it into replicate disposable test tubes or PE transfer pipettes. 8. Wrap Parafilm ~ tightly a r o u n d the m o u t h of each test tube and heatseal the stem of each PE transfer pipette. 9. Place the inoculated p o u r tubes in a pressure vessel that is thermally equilibrated to the desired incubation temperature and compress it to the desired pressure. Dissipation of the adiabatic heating due to compression results in a small and negligible pressure drift with aqueous hydraulic fluids. 10. In the example here, the final p H of the gel should be 7.8. To obtain this p H for unbuffered sols or a different p H for buffered sols, an empirical determination of the n e e d e d concentration of N a O H is m a d e with a test batch of silicic acid sol. A drawing of a PE bulb with colonies of bacteria is s h o w n in Figure 30.4.
Doryaki method N a k a y a m a et al. (1994) prepare two sterile slabs of nutrient agar. They spread cells or streak them onto one of the agar slabs and then place the other slab over it to form an agar sandwich or 'doryaki'. They place the agar 'doryaki' in a plastic pouch, heat-seal it, and incubate it at high pressure. It w o u l d be of interest to see if motile strains form either discrete colonies or a diffusely growing mass of cells w h e n placed between two outer slabs and incubated in a pressure vessel.
ENRICHMENT PRESSURE
CULTURES AT HIGH
The heat-sealable PE bulbs described for the colony-forming ability assay also allow for a variety of enrichment cultures. For example, sulfatereducing bacteria from sea floor sediments collected at a water d e p t h of 3600 m grew in these bulbs. The heat-sealed, inoculated PE bulbs were in an oxygen-impermeable bag filled with a dithionite solution that scavenges dissolved oxygen. The bacteria grew during the incubation of the bag in a pressure vessel at ca. 36 MPa and remained anoxic.
MAINTENANCE
OF P I E Z O P H I L I C B A C T E R I A
Many bacteria from the cold deep sea survive storage in the presence of a cryoprotectant at -80°C. Cells can also be maintained in culture m e d i u m
631
at high pressure and 2°C with periodic transfer to fresh medium. Lyophilization failed repeatedly as a means of preservation for two deepsea bacterial strains and we chose not to pursue the possibility of this mode of storage any further.
H I G H PRESSURE L A B O R A T O R Y I N S T R U M E N T S FOR G R O W T H CHARACTERIZATION The above techniques allow for the isolation of pressure-adapted microorganisms and for determination of the lethal effects (that is, loss of colonyforming ability) of stress. The study of pure cultures requires instruments for determining growth rates and other physiological processes. Three instruments are given a brief description here.
Kinetics of microbial growth: sampling with decompression The techniques described in previous sections of this chapter show how to incubate liquid cultures in bags or syringes containing glass or Teflon balls. The culture containers are in pressure vessels that are on a rocking platform in a temperature regulated water bath. We sample a culture in a pressure vessel by decompressing it, removing the culture container, removing a portion of the culture, returning the culture container to the pressure vessel, and restoring the pressure. With the use of pin-closure vessels and the quick-connect fitting, the sampling procedure can be done within two minutes. We call this process sampling with decompression/compression cycles. The physiological effects of a two-minute cycle of decompression and compression require further study to establish general confidence in the results. Nevertheless, we are moderately certain that growth rates of bacteria with long generation times are accurate when determined by this method. However, when the interval of decompression for sampling is, for example, 5 or 10% of the generation time of the organism, then the growth rates may be inaccurate.
Kinetics of microbial growth: sampling without
decompression
An ideal high-pressure kinetics system is one that allows removal of a sample from a culture in a pressure vessel while avoiding both decompression of the bulk of the culture and exposure of the sample to any shear forces. Designs for such instruments are given in several publications (Yayanos, 1969; Jannasch et al., 1973; Jannasch et al., 1976; Kelly and Deming, 1988; Yayanos et al., 1983; Yayanos, 1998; Jannasch et al., 1996). This is not a complete list of efforts to design high-pressure kinetics systems: inspection of these papers, however, reveals that these methods are largely of a prototype nature.
632
PRESSURIZED T E M P E R A T U R E G R A D I E N T INCUBATORS Polythermostats allow the incubation of organisms at a large number of temperatures along a temperature gradient (Thomas et al., 1963). A pressurized temperature gradient (PTG) is an incubator that provides the polythermostat capability at several different high pressures (Yayanos et al., 1984; Sakiyama and Ohwada, 1997). Incubation of bacterial cultures in a PTG provides data on the approximate location of the pressure temperature boundary within which the organism can grow. Incubations can also give an idea of the approximate temperature and pressure at which the cells grow fastest. The full potential of these instruments in microbiology remains to be explored.
t,eee4,
SAFETY ISSUESWITH HIGH-PRESSURE EQUIPMENT There is no single source of safety information for the conduct of highpressure research. Comings (1956) devotes an entire chapter, albeit short, to safety. The following is an incomplete minimal list of safety issues to keep in mind. Each investigator in high-pressure research should take time to analyze the possible modes of failure in their apparatus and of other hazards that may be specific to their experimental design. 1. Wear safety glasses or, preferably, a clear plastic face cover when operating high-pressure equipment. 2. Do not exceed the working pressure limit of the weakest component in your high-pressure system. 3. Use a rupture disk assembly that is always online when you are compressing the pressure vessel. Select a rupture disk that will fail at a pressure well below the test pressure of the weakest component in your system. 4. Determine whether any unusual chemical reactions might occur in your experimental system. For example, do not use ammonia in contact with liquid mercury because compounds form at high pressures that detonate on decompression (Comings, 1956). 5. If you must use hydrogen gas or carbon monoxide, work with scientists, engineers, or companies who are familiar with the potential problems. 6. Decompress a vessel through a valve and tube so that the fluid expelled discharges into a waste reservoir. 7. Tubing having an outer diameter of ~ inch, although conveniently flexible, will become weak from repetitive flexing. It is good idea to replace such tubing periodically or at any indications of weakness. High-pressure liquid discharged through ~ inch fittings can cut skin, damage eyes, and inject microorganisms into personnel. 8. The amount of energy released when a component of a high-pressure system fails depends on the compressibility and volume of the 633
compressed material. Thus, compressed gases present far greater hazards than do liquids. Large pressure vessels u n d e r pressure are more hazardous than small ones. Where possible, use barricades between the experimentalist and the equipment. 9. Make sure that the sample container inside a pressure vessel cannot block the hole that leads to the pressure relief valve. If this hole becomes blocked, then it is possible that the pressure inside the vessel will rupture the sample container w h e n you open the pressure relief valve. If the pressure relief valve is not connected to a tube leading to a waste reservoir, then an aerosol of microorganisms will be formed. 10. Insure that the temperature of a water bath or incubator housing a sealed pressure vessel cannot go too high since the thermal expansion of the vessel contents will cause a rise in pressure. Either prevent such a temperature increase from occurring or ascertain by calculation that the pressure inside of the closed pressure vessel will not exceed the u p p e r working pressure for that vessel. 11. When using a pin-closure vessel, ascertain that the pin is properly in place before increasing the pressure. A good practice is to keep the pin in place with a rubber band or clamp. N e v e r compress a pin-closure vessel without the pin in place. This could lead to the cap becoming a projectile. Never point the cap of a pressure vessel towards you or anyone else. 12. Never look directly into the w i n d o w of a pressure vessel. Even viewing via a mirror can be unsafe because of the possibility of reflected material during a failure. 13. Before adopting the design of a high-pressure device from a scientific publication, review it carefully to authenticate its design limits and identify possible printing errors. Scientists often use an apparatus for demonstration purposes. If the apparatus were to be sold commercially, then the design w o u l d likely be modified to reflect safety codes.
Acknowledgements I thank the National Science Foundation for financial support of my research under grant OCE9907651.
References Berger, L. R. and Tam, L. Q. (1970). A method to grow obligately aerobic bacteria at increased hydrostatic pressure. Limnol. Oceanogr. 15, 483-485. Bianchi, A., Garcin, J. and Tholosan, O. (1999). A high-pressure serial sampler to measure microbial activity in the deep sea. Deep-Sea Res. Pt. I. -Oceanog. Res. 46, 2129-2142. Comings, E. W. (1956). High Pressure Technology. McGraw-Hill, New York. Deming, J. W. (1997). Unusual or extreme high-pressure marine environments. In: Mammal of Environmental Microbiology (C. J. Hurst, Ed.), pp. 366-380. American Society for Microbiology. Deming, J. W. and Baross, J. A. (1986). Solid medium for culturing black smoker bacteria at temperatures to 120°C. Appl. Environ. Microbiol. 51, 238-243.
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Dietz, A. S. and Yayanos, A. A. (1978). Silica gel media for isolating and studying bacteria under hydrostatic pressure. Appl. EnviroJ~. Microbiol. 36, 966-968. Dodge, B. E (1950). High pressure technique, in: Chemical Etl~imvrs Ha~ldbook (J. H. Perry, Ed.), pp. 1233 1262. McGraw Hill, New York. Fryer, D. M. and Harvey, J. E 11998). tti~¢h Pressure Vessels. Chapman & Hall, New York. Gasche, E 11966) Pressure Vessel. USA Patent [3,258,151 ]. Hawley, S. A. 11978). High-pressure techniques. Methods EHzymol. 49, 14-24. t telmke, E. (1979). Zur Kultivierung yon aeroben ActiJtomycete11 mariner und terrestrischer Herkunft unter hydrostatischenl Druck. Vcr{!ff. hist. Meeres[orsch. Brt'mcdl. 18, 1-20. Holzapfel, W. B. and Isaacs, N. S. (1997). High Press~trc TechHiques itl Chemistry amt Physics: A Practical Approach. Oxford University Press: Oxford. Horikoshi, K. and Tsujii, K., eds (1999). Extremophih's ill Deep-Sea Enviromm'uts. Springer-Verlag, Tokyo. Jannasch, H. W. and Wirsen, C. O. 11984). Variability of pressure adaptation in deep sea bacteria. Arch. Microbio[. 139, 281-288. Jannasch, H. W., Wirsen, C. O. and Winget, C. L. (1973). A bacteriological pressureretaining deep-sea sampler and culture vessel. Deep-Sea Res. 20, 661-664. Jannasch, H. W., Wirsen, C. O. and Taylor, C. D. 11976). Undecompressed microbial populations from the deep sea. Appl. EJlviroJt. Microbiol. 32, 360-367. Jannasch, H. W., Wirsen, C. O. and Taylor, C. D. 11982). Deep-sea bacteria: Isolation in the absence of decompression. Scieltce 216, 1315 1317. Jannasch, I t. W., Wirsen, C. O. and Dohert)5 K. W. (1996). A pressurized chemostat for the study of marine barophilic and oligotrophic bacteria. Appl. E~i~,irom Microbiol. 62, 1593 1596. Johnson, E H., Eyring, H. and Polissar, M. J. (1954). The Killetic Basis of Molecldar Biology, pp. 286 368. John Wiley & Sons, Inc., New York. KelI> R. M. and Deming, J. W. 11988). Extremely thermophilic archaebacteria: Biological and engineering considerations. Biotechnol. Prog. 4, 47-62. Kluyver, A. J. and van Niel, C. B. (1956). Tht' Microbe's Co~#ributioJ7 to Biology. Harvard University Press, Cambridge, MA. Ludlox~, J. M. and Clark, D. S. 11991). Engineering considerations for the application of extremophiles in biotechnology. Crit. Roy. Biotcctmol. 10, 321-345. Macdonald, A. G. 11978). Further studies on the pressure tolerance of deep-sea crustacea, with observations using a new high-pressure trap. Mar. Biol. 45, 9-21. Macdonald, A. G. and Gilchrist, I. (1969). Recovery of deep seawater at constant pressure. Nature 222, 71-72. Marteinsson, V. T., Birrien, J. L., Reysenbach, A. k., Vernet, M., Marie, D., Gambacorta, A., Messner, P., Sleytr, U. B. and Prieur, D. (1999). Thermococcus barot~hihis sp. nov., a new barophilic and hyperthermophilic archaeun isolated under high hydrostatic pressure from a deep-sea hydrothermal vent. hH. J. Syst. Boot. 49, 351-359. Murray, C. N., Stanners, D. A. and Jamet, M. 11989). Technical note: A piston corer for recovery of deep ocean sediments nnder pressure. Mar. Gcoteclmol. 8, 69-80. Nakayama, A., Yano, Y. and Yoshida, K. 11994). New method for isolating barophiles from intestinal contents of deep-sea fishes retrieved from the abyssal zone. Appl. Eplvirou. Microbiol. 60, 4210-4212. Newitt, D. M. 1194/)). The Design of High Pressure l)la1# alat the lhopertics of Fluids at High Pressures. Oxford University Press, Oxford. Sakiyama, T. and Ohwada, K. (1997). Isolation and growth characteristics of deepsea barophilic bacteria from the Japan trench. Fisheries Sci. (Tokyo) 63, 228-232. Suzuki, K. (1973). Measurements at high pressure. Meth. Enzymol. 26, 424-452.
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Tabor, P. S., Deming, J. D., Ohwada, K., Davis, H., Waxman, M. and Colwell, R. R. (1981). A pressure-retaining deep ocean sampler and transfer system for measurement of microbial activity in the deep sea. Microbial. Ecol. 7, 5i-65. Thomas, W. H., Scotten, H. L. and Bradshaw, J. S. (1963). Thermal gradient incubators for small aquatic organisms. Limnol. Occm~ogr. 8, 357-360. Tongue, H. (1959). The design and construction of high pressure chemical plant. Van Nostrand: Princeton, NJ. Tsiklis, D. S. (1968). In: Handbook q~ Techniques in ttigh-Pressurc Research a~uf En~,ineeriny, (A.T. Bobrowsky, Ed.). Plenum Press, New York. Vodar, B. and Saurel, J. (1963). The properties of compressed gases. In: High Pressure Physics a,d Chemistry (R. S. Bradley, Ed.), pp. 51-143. Academic Press Inc., New York. Yayanos, A. A. (1969). A technique for studying biological reaction rates at high pressure. Rev. Sci. lnstrum. 40, 961-963. Yayanos, A. A. (1978). Recovery and maintenance of live amphipods at a pressure of 580 bars from an ocean depth of 5700 meters. Science 200, 1056-1059. Yayanos, A. A. (1980). Measurement and instrument needs identified in a case history of deep-sea amphipod research. In: Advanced Concepts i, Ocea, Measun'ments fi~r Marine Biology (F. D. Diemer, E J. Vernberg, and D. Z. Mirkes, Eds), pp. 307-318. University of South Carolina Press, Columbia, SC. Yayanos, A. A. (1982). Deep sea biophysics. In: Subseabed Disposal Program Ainu,71 Report ]amlary to September 1981. Volume 11: Appendices. Part 2 of 2 (K. R. Hinga, Ed.), pp. 407-426. Sandia National Laboratories, Albuquerque, NM. Yayanos, A. A. (1995). Microbiology to 10,500 meters in the deep sea. Amt. Rev. Microbiol. 49, 777-805. Yayanos, A. A. (1998). Empirical and theoretical aspects of life at high pressures in the deep sea. In: Extremophiles (K. Horikoshi and W. D. Grant, Eds), pp. 47-92. John Wiley & Sons, New York. Yayanos, A. A. (in press). ZoBell and his contributions to piezobiology (barobiology). In: Microbial Biosystems: New Frontiers. Proceedings of the 8th Eighth h#erluTtional Symposium ot~ Microbial Ecology (C. R. Bell, M. Brylinsky, and 12 Johnson-Green, Eds). Atlantic Canada Society for Microbial Ecology, Halifax, Canada. Yayanos, A. A. and DeLong, E. E (1987). Deep-sea bacterial fitness to environmental temperatures and pressures. In: Current Perspectives i~ High Pressure Biology (H. W. Jannasch, R. E. Marquis and A. M. Zimmerman, Eds), pp. 17-32. Academic Press, New York. Yayanos, A. A. and Dietz, A. S. (1983). Death of a hadal deep-sea bacterium after decompression. ScieHce 220, 497 498. Yayanos, A. A. and Van Boxtel, R. (1982). Coupling device for quick high-pressure connections to 100 MPa. Rev. Sci. lnstrum. 53, 704-705. Yayanos, A. A., Van Boxtel, R. and Dietz, A. S. (1983). Reproduction of Bacillus stearothermophilus as a function of temperature and pressure. Appl. Em,irotl. Microbiol. 46, 1357-1363. Yayanos, A. A., Van Boxtel, R. and Dietz, A. S. (1984). High-pressure-temperature gradient instrument: use for determining the temperature and pressure limits of bacterial growth. Appl. Emqron. Microbiol. 48, 771-776. ZoBelI, C. E. (1952). Bacterial life at the bottom of the Philippine Trench. ScieHcc 115, 507-508.
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List of suppliers Fisher Scientific 2761 Wahlut Avenue Tustin, CA 92780, LISA TH: 1-800-766-7000 Fax: 1-800-926-1166 http://www3.fishersci.com/dhtlnl./sp
Chemicals, Samco polyethylene transfer pipettes, heat sealers Harwood Engineering Company, Inc. 455 South Shvet Walpole, MA 02081, USA Teh 508-668-3600 Fax: 508-660-2276 http://www.tdtranet.com/~harwood/ Emaih [email protected]
Specializes in equipment for very high pressures
High Pressure Equipment Company P.O. Box 8248, 1222 Linden Avenue Erie, PA 16505, USA TH: (814) 838 2028 1 80(]-289-7447 Fax: (814) 838-6075 httl~://www.hight~ressmv.conl/
High pressure equipment, fittings, valves, adapters, gauges, pumps, pressure vessels Pressure Products Industries, Inc. 900 Louis Drive, Wanninster PA 18974, LISA Teh (215) 675 1600 Fax: (215) 443-8341
iJlfo@,~otolpi.com High pressure equipment
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1050 Arroyo Avemte Sail Fernando, CA 91340, LISA Teh 818/838-2400 Fax: 818/838 2488 E HIaH:cIIstFscr[[email protected]
High pressure equipment, fittings, valves, adapters, gauges, pumps, pressure vessels, and air-driven high pressure pumps
Polylethylene transfer pipettes Sarstedt, Inc. 1025 St. James Church Road PO Box 468 Newton, NC 28658, LISA Teh 800/465-4000 Fax: 828/465-0718
Disposable plastic syringes
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31 Methods for the Study of Hydrothermal Vent Microbes Anna-Louise Reysenbach and Dorothee G6tz Deportment of Biology, 1719 SW I Oth Avenue, Portland Sto1:eUniversity, Portlond, OR 9720 I, USA
CONTENTS Introduction Sampling procedures Shipboard sample handling Methods for estimating diversity Future developments
4,ee~,ee INTRODUCTION With the discovery of deep-sea hydrothermal vents in 1979 came the discovery of unique invertebrate communities associated with these very unusual environments. Much of the initial biological research at deep-sea vents focused on new descriptions of the invertebrates, although the central role that microbes play in the deep-sea h y d r o t h e r m a l vent ecosystem was realized early on. This initial research explored the role of symbionts in their invertebrate hosts, and the activities of the free-living sulfur-oxidizing mesophilic microbial communities. Furthermore, with the discovery of deep-sea vents, and hydrothermal fluids at temperatures that exceed 350°C, the possibility that life could exist above 100°C was realized. Consequently there was an increased interest in the diversity, ecology and physiology of thermophilic microorganisms. An excellent review of microbial research at deep-sea vents is available in the book edited by Karl (1995). In this chapter we explore ways in which microbiological samples can be obtained from deep-sea vents and h o w samples m a y be processed for DNA extraction or enrichment culturing. This is not an exhaustive list of m e t h o d s and e q u i p m e n t required for deep-sea microbiological research, as in m a n y cases specialized instruments have to be designed for specific research purposes. Additional web pages and resources are listed at the end of the chapter.
METHODSIN MICROBIOLOGY,VOLUME30 ISBN 0-12-32153(I~4
Copyright © 2001 Academic Press Ltd All rights of reproduction in anv form reser~ed
•
SAMPLING
PROCEDURES
Perhaps one of the most challenging aspects of doing deep-sea microbiological research is obtaining a representative and uncontaminated sample. The types of samples that can be collected include high-temperature fluids, diffuse flow (low-temperature, shimmering water) water samples, microbial mats, rock and sulfide mineral samples, epibionts and symbionts associated with animals.
Sampling vehicles Numerous research submersibles are available. In the United States, the Woods Hole Oceanographic Institution (WHOI) manages the deep-sea submersible, DSV Alvin and the remotely operated vehicle, DRV Jasom The specifications for these vehicles can be obtained from the WHOI webpage (www.whoi.edu). Requests for use of the WHOI-operated vehicles as part of nationally funded scientific research are made to tile University National Oceanographic Laboratory System (UNOLS) (http://www.gso.uri.edu/unols/unols.html). Other deep-sea research submersibles include the Ifremer (Brest, France) operated vehicle, DSV Nautile, the Russian operated vehicles, DSV MIR, and the Japanese vehicles, DSV Shinkai 2000 and 6500. Special precautions should be taken if specific equipment is to be used on the submersibles. For example, all equipment that is sealed or has an external housing, needs to be pressure certified. If the piece of research equipment needs to use the submersible power, specific arrangements should be made with the submersible staff and the researcher as to the compatibility of equipment with the submersible. The different submersibles have different configurations of robotic arms, and how submersibles pick up and trigger an instrument will differ considerably depending on the configuration of the submersible hydraulic arms. Although the Ocean Drilling Program (ODP) research vessel (RV loides Resolution) has not been considered a vessel for routine microbiological sampling, recent interest in exploration of the extent of the deep subsurface biosphere has stimulated the establishment of a microbiological laboratory on the ship. Scheduled equipment includes a fluorescent microscope, pressure vessels and culturing facilities.
Elevators versus baskets The submersibles and remotely operated vehicles (ROVs) are equipped with a large basket onto which sampling containers and equipment can be attached and into which all samples collected during a research dive are placed. However, space on the basket is often very limited, and some submersible operations use an 'elevator' (a basket attached to a cable) to take more cumbersome equipment down to the seafloor. The submersible or ROV uses the elevator to pick up and drop off equipment and large 640
samples. The D S V Nautile operation uses an elevator routinely, as do some of the ROV teams.
Pressure considerations Deep-sea hydrothermal vents have been explored to depths of over 3700 m. Pressure therefore is a consideration when exploring a microbial sampling strategy. Several laboratories (including Japan Marine Science and Technology Center (JAMSTEC)) have designed microbial sampling devices that maintain the in situ pressure; to our knowledge none are used routinely. A description of one such device that is also an in situ growth experiment is described below. Less emphasis has been placed on retrieving samples under pressure, as numerous studies have shown that many of the isolates that have been isolated from deep-sea vents are barotolerant. Two isolates, Thermococcus barophilus (Marteinsson ct al., 1999) and Palaeococcus fertvphilus (Takai et al., 2000) do appear to be moderate barophiles or piezophiles. For more details on barophilic bacteria, see Chapter 30 of this volume.
In situ sampling devices
Water samples A number of different water sampling devices have been developed primarily for geochemical analyses. The titanium (Ti) syringes or 'major samplers' (Von Damm et al., 1985a,b) can sample up to 755ml of hydrothermal fluid. These syringes are not gas-tight. If gas samples are required, the 'Lupton sampler' (Lupton et al., 1985, Lilley et al., 1994) can be used for exclusive gas measurements. Recently, Jeff Seewald at WHOI, developed a gas sampler in which the sample is maintained at seafloor pressure during removal of the fluid onboard ship. Subsamples can be removed for different chemical analyses in addition to gas analysis. Additionally, the controlled fill rate (about 2 rain for a 150 ml sample) makes the sampler ideal for diffuse venting. Gas concentrations such as t-L and CO, are fundamental requirements of many chemolithotrophs; therefore obtaining gas measurements, in addition to standard geochemical measurements, are important for understanding the ecological niche of these thermophiles. A variation of these samplers is the submersiblecoupled, ilz situ sensing and sampling system, SIS ~, which can accommodate three Ti or gas-tight samplers. This system allows for the samplers to be flushed before an actual sample is taken, and the temperature at the inlet nozzle and at the intake of the last sampler in the series can be obtained. A Hydrothermal Fluid Particulate Sampler (HFPS) was recently designed at National Oceanic and Atmospheric Administration - Pacific Marine Environmental Laboratory (NOAA-PMEL) (Figure 31.1) which is a significant advance over many of the in situ samplers. This system allows for up to 24 independent samples to be taken and can measure the
641
Figure 31.1. Hydrothermal Fluid Particulate Sampler (HFPS). In tile front, from left to right, the 25-port valve, the flushing pump, the sample pump, the power and control unit. Filters are visible across the middle of the apparatus, and with piston samplers and 8 bag samplers towards the back. Filters can be placed in front of bags. The sampling inlet tool is on the right. (Courtesy David Butterfield, NOAA-PMEL.)
temperature where the sample is being taken. It has additional advantages for microbial sampling over other systems in that in-line filters can be used to concentrate cells and biological fixatives m a y be used. This latter application will provide snapshot sampling, but also allows for stabilizing DNA and RNA for m R N A measurements. The 'OsmoSampler' is a device for continuous fluid sampling over periods from days to months (Jannasch et al., 1994). It has primarily been used for geochemical sampling, however, if it were to be coupled to microbiological sampling, it m a y prove to be a very valuable deep-sea microbial monitoring tool. N u m e r o u s other sampling devices have been designed. An 'off the shelf' rosette of syringes (6-12) for water sampling has been designed by General Oceanics Inc. In this case, the intake tube can be m o u n t e d in a 'wand' that also has a temperature-sensing device and in this way the submersible's hydraulic arm can position the intake tube where desired (Figure 31.2). A pelagic p u m p is placed in the line and water can be p u m p e d through the w a n d and into the syringes. The limitation of this type of system is the void v o l u m e that the sampling lines take u p and the u p p e r temperature that the wand material can tolerate. The buoyant hydrothermal plume or surrounding seawater can be sampled by using Niskin bottles that are adapted for triggering by the submersible and are attached to the submersible basket. The volumes can vary from 1.5 to 5 1. The problem with Niskin bottles is that there is little or no control for possible entrainment of surrounding seawater. A more 642
(~)
(b)
Figure 31.2. (a) Rosette Syringe Sampler designed by General Oceanics Inc. (b) the fluid intake 'wand' taking a fluid syringe sample at a deep-sea hydrothermal chimney on the East Pacific Rise. (Courtesv Luther, Reysenbach and Cary.)
effective device perhaps for microbiological research of hydrothermal plumes and diffuse flow, has recently been developed by G e r m a n et al. (in preparation) and is called the Buoyant Plume Sampler (BPS). This is a very compact, light instrument for performing iH situ filtration and is easily used by a range of ROVs and m a n n e d submersibles.
Microbial mat and sediment samples
H y d r o t h e r m a l l y active areas such as tile G u a y m a s Basin also contain hot sediments that are covered by thick microbial mats. Sediment and mat samples can be obtained using box corers or regular push corers. Again, the corers need to be able to withstand temperatures that may exceed 100°C. Degassing of the cores during ascent to the surface can cause turbulence in the cores that disrupts core stratification. Surface mats can be collected using a firm inlet pipe (regular v a c u m n cleaner hose works!), connected to a pelagic p u m p and sampling bottles. This device is sometimes referred to as a 'slurp gun'. The inlet to the sampling bottles m a y have filters of desired mesh size to prevent certain sized objects (such as rock particles) from being collected and contaminating the p u m p .
Rocks, animals and active sulfide structures
Animal or rock samples can be collected by the submersible arm and placed in an insulated container (bio-box), in the submersible basket. Although not completely sealed, a container will minimize flushing and contamination of samples bv s u r r o u n d i n g seawater as the submersible 643
returns to the ship. In addition, the insulated box will keep the samples cool, and prevent them from heating up in the warm surface waters.
In situ growth and activity measurements Numerous in situ samplers have been designed in an attempt to obtain microbial samples of actively growing cells and for analyzing in situ microbial activity. The simplest approach has been to use the concept of 'contact slides' or 'Cholodny-Rossi slides' (Atlas and Bartha, 1993). For example, any instrument placed near hydrothermal vents is covered very rapidly with microbial growth (Taylor et al. 1999). Using this rationale, Norman Pace and David Lane (see Karl et al., 1988) designed the 'vent cap' (Figure 31.3), a titanium cylinder with a cone-shaped base. Different surfaces for microbial attachment are inserted in the cylindrical chamber. The 'vent cap' can be placed on top of an active hydrothermal chimney; the hydrothermal fluid passes continuously over the surfaces providing nutrients for microbes that happen to attach and proliferate on the surfaces. The instrument can be deployed for a desired period of time (1 to 5 days is recommended). The chamber is closed and brought to the surface where the 'vent cap' surfaces are analyzed by microscope and serve as inocula for enrichment culturing or are analyzed using molecular phylogenetic approaches. The design has been modified from its original spring-loaded design to incorporate a simple slide mechanism for closing the chamber and inclusion of a datalogger and temperature probe for continuous monitoring of the temperature within the chamber. This instrument could also be used to study microbial colonization and succession at hydrothermal vents. The communities that colonize the surfaces in the 'vent cap' are probably a subset of the true microbial diversity found at vents, since only organisms capable of attaching to the surfaces provided will proliferate. An even more selective approach to obtaining microbial samples at vents is the use of 'biotraps'. These were first used by the company DIVERSA, to concentrate and select for certain thermophiles from hot springs in Yellowstone National Park. DIVERSA and others have subsequently used them at deep-sea hydrothermal vents. These units are packages of porous spheres that are impregnated with organic substrates for specific enrichment of organotrophs. The 'biotraps' are deployed for days to months at predetermined locations. The beads then serve as a source of inoculum or DNA. Other less biased designs have been used successfully, such as the 'Bioblanket ' and Biocolumn' designed by Jim Cowen and Paul Johnson at the University of Hawaii and University of Washington, respectively. Microelectrodes have been used successful for inferring microbial processes in hydrothermal sediments (J•rgensen et al., 1990) and measuring bioavailable redox species using voltammetric microelectrodes (Luther et al., 1998). However, f e w instruments have been designed to measure microbial activity in situ, in particular at high temperatures. One such instrument, the 'Laredo' sampler, was developed by Deming and others at the University of Washington and lfremer, France. This instrument is also placed on hydrothermal chimneys, and once the fluid
644
(at
(b)
¢-
..0
0
Figure 31.3. (a) Diagram of the titanium 'vent cap' or in situ growth chamber. A is the datalogger that records tile temperature within B, the chamber. Surfaces are placed in B and once the slide C is opened, fluid can be channeled through D, tile cone-shaped skirt, over tile surfaces and out through the base of B. (b) The 'vent cap' being deployed on a hydrothermal vent in Guaymas Basin off Mexico. The blurriness is the hot 'shimmering' hydrothermal fluid. The white floc in the water column is from bacterial mats that have been dislodged. The instruments in the foreground are push corers for taking sediment and mat samples.
645
"0 >,,
I
flowing through the chamber reaches a specific temperature, the chamber is triggered to close by a microcomputer attached to the instrument package. A radioactive substrate in a glass vial may be placed in the chamber, which can be released to start a specific in situ activity experiment. Once complete, the sampler can be returned to the surface and the contents of the sealed chamber analyzed. The sampler has recently been deployed successfully on hydrothermal vents along the East Pacific Rise. Clearly, these types of experiments are very costly and difficult to accomplish, due to the many different problems that may arise in a complex set up. Furthermore, they represent one single time point, and many successful deployments are required to provide a statistically relevant sample size. However, deep-sea microbiologists are not alone in the limitation of sample size. Macroecology in the deep-sea is also often criticized due to lack of statistically relevant sampling. This is one caveat of deep-sea biological research that we will have to learn to accept and try to address more rigorously.
4,4,4,4,4,4, S H I P B O A R D S A M P L E H A N D L I N G Once shipboard, samples should be processed immediately. Subsamples from water bottles can be taken using sterile syringes and used as inoculum for enrichment media. Animals should be kept on ice, and epibionts can be removed using sterile tweezers or spatulas. Sediment cores should be extruded immediately, and can be processed in a disposable anaerobic glove bag if necessary. Sulfides should be sectioned as soon as possible. Using a saw with a tungsten carbide blade (normally used to cut ceramic) is recommended to section hard sulfides. Many sulfides are very fragile and can be sectioned with sterile spatulas and subsampled. Sulfide samples can be ground in a disposable anaerobic glove bag with a sterile pestle and mortar. The samples are normally moist enough to not require any addition of liquid. The ground sample can be inoculated into anaerobic media. If the DNA in the sample needs to be stabilized, grinding the sample in a DNA buffer is recommended. For some hyperthermophiles, such as the acidophiles, the pH of the sample may need to be adjusted above pH 6. The internal pH of these acidophiles rapidly equilibrates to the surrounding pH if they are not growing, causing cell lysis. Since samples are rapidly cooled once collected from deep-sea vents, it may be too late to wait till they reach the surface to neutralize the samples. A possible reason why acidophiles such as Su(fi~lobus have not been isolated from deep-sea vents is that the pH of samples has not been stabilized in situ. Additionally, Su(folobus is a strict aerobe, and stable conditions where the oxygen levels are high and temperatures are above 70°C may not exist at deep-sea vents.
Useful equipment to have shipboard A n example of a packing list for a deep-sea oceanographic cruise can be obtained at http://caddis.esr.pdx.edu/alr
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• • • • • • • • • • • •
eeeeee Molecular
Digital thermometer A hand saw with a tungsten carbide blade (essential for cutting sulfides) Tubing Tools (large wrench for gas tank) Gas manifold Coring devices Containers for sharp disposal Trays for placing samples on Lots of culture tube racks General lab supplies Autoclave Microscope Having access to a microscope with phase contrast and fluorescence capabilities is highly recommended. Although looking at microbes under high magnification at sea can be tiring and stimulate nausea, the advantages far outweigh this temporary discomfort.
METHODS
FOR ESTIMATING DIVERSITY
methods
D N A extraction and molecular biological approaches
MJ scientific makes a 'Mobile Molecular Laboratory' (Model MML-0150) that is compact and easily transported. It is r e c o m m e n d e d that prep o u r e d gels be used for shipboard electrophoresis. Procedures that minimize the use of hazardous chemicals such as phenol are recommended. Most general DNA extraction procedures are adequate for samples collected at deep-sea vents. However, sulfides and sediments that are rich in metals are problematic for DNA extraction as the metals bind to the DNA and can inhibit the PCR. Below is one procedure that has worked reasonably well in our laboratory, and manipulating extraction protocols with elevated chelators such as EDTA and the addition of ion-exchange resins such as Chelex 100 (Sigma C7901) does appear often to increase DNA yields. DNA extraction from bacteria associated with sulfide minerals protocol (after Dempster eta/., 1999). 1. Suspend sample in 500 ~tl of pre-heated extraction buffer: 100 mM Tris-Cl (pH 8.0) 1.4 M NaC1 20 mM EDTA ().4~/~ (vol/vol) 2-mercaptoethanol 2 ~ ( w t / v o l ) c e t y l t r i m e t h y l a m m o n i u m (CTAB) (Aldrich 855852) 1 ~7( p o l y v i n y l p y r r o l i d o n e (PVP 360, Sigma). 2. Incubate for 15 rain at 65~'C. 3. Add 500 ~tl of chloroform:isoamyl alcohol (24:1) and w)rtex for 2 rain. 4. Spin for 15 rain and transfer aqueous layer to a fresh tube. 647
5. Precipitate DNA with an equal volume of isopropanol and 0.5 volume of 5 M NaC1 at room temperature for 15 min. 6. Spin for 30 min and wash with 70G ethanol. 7. Re-suspend in 100 ~l of TE and add 1 ~tl of RNAse (1 mg ml '). 8. Incubate at 37°C for 30 min. 9. Precipitate DNA with 2 volumes of freezer-cold ethanol and 0.1 volume of 3 M sodium acetate at 4°C for 1 h. 10. Spin for 30 rain at 4°C, wash with freezer-cold ethanol, dry pellet. Including steps using guanidine thiocyanate and silica beads may increase yields in some samples (Huang et al., 2000).
Enrichment, culturing, isolation and maintenance An excellent resource for enrichment culturing media for thermophiles is R o b b c t al. (1995) or the webpage for Deutsche Stammsammlung yon Mikroorganismen und Zellkulturen (http://www.dsmz.de/). These media are a good base for enrichment of specific physiological types. If your interest is in growing the diversity of metabolic types that occur at deep-sea vents, using classical, well-designed and proven media recipes may not achieve this objective. It is likely that similar isolates will be obtained from geographically diverse areas. Taking a more ecological approach will no doubt lead to new isolates that have never been isolated from deep-sea vents (for example, Reysenbach et al., 2000). Determining the environmental conditions in which these organisms exist will provide clues to the energy and carbon sources available to growth. In situ geochemical measurements using microelectrodes provide measurements at a biological relevant scale. Using geochemical modeling, one can begin to predict which redox reactions are energetically favorable (McCollom and Shock, 1997), and this will help develop novel enrichment media. Parameters such as temperature, pH, chemical composition, salinity, pressure and oxygen, hydrogen, carbon dioxide and other gases are guidelines for media composition and incubation conditions. Microorganisms vary in their ability to adapt to suboptimal conditions; it is therefore best to mimic the enviromnental setting as closely as possible. Additionally, at deep-sea vents where fluctuating chemical and physical gradients are the norm, strategies that explore these gradients and the anoxic and oxic interfaces may reveal a novel culturable diversity not previously explored by deep-sea microbiologists. The choice of electron acceptors, electron donors and carbon sources supplied in the culture medium will help enrich for specific metabolic types. If the aim is to isolate as wide a physiological diversity from one habitat as possible then a matrix of electron acceptors, electron donors, carbon sources, temperatures, pH, salt concentrations and concentrations of oxygen should be designed. Because of the ease with which microbes can be identified by molecular phylogenetic techniques, we routinely assess the molecular phylogenetic diversity of our initial enrichments. These initial enrichments are often very diverse, but already very different from the original diversity of the
648
sample from which they were enriched. Using this phylogenetic information, we are then able to follow and select for organisms that are of interest to a project. Furthermore, in liquid media, slow-growing organisms are frequently overgrown by faster growing ones and are often overlooked and lost. This combination of enrichment culturing guided by molecular phylogenetic identification early on in the enrichment culturing process helps overcome some of these problems. Additionally, some organisms can be isolated more easily by direct plating of the sample on plates or in roll tubes. Some microbes from deep-sea hydrothermal environments might require high pressure for growth. We will not explore these conditions; see Chapter 30, this volume, for the isolation and cultivation of barophiles, and Baross and Deming (1995).
Medium preparation A very useful compilation of media for tile cultivation of thermophiles isolated from marine hydrothermal vents can be found in Robb ct al. (1995, Appendix 2). A basic enrichment medimn can be formulated based on the solutions detailed below. This basic recipe can then be adapted with the addition of the various electron donors, electron acceptors and carbon sources, suitable nitrogen, sulfur, phosphorus sources, nutrients and in some cases vitamins, and appropriate pH and incubation temperature.
Culturing equipment: • Bellco tubes and flasks • Serum bottles • Gas impermeable butyl rubber stoppers • Aluminum crimp seals, crimper • Plastic or glass Petri dishes • Chemicals (salts, organics, vitamins) • Gases and gas mixtures (N2, 02, H~, CO2, CO2-H~, CO2-N2) • Gas manifold • Incubators • Shaking incubators • Needles and syringes
e-
Base solutions for the e n r i c h m e n t m e d i a
b
Artificial (synthetic) seawater (in g 1 '): NaCI 20.0 g MgCI~-6H~O 3.0 g MgSO,.7H~O 6.0 g (NH,).SO; 1.0 g NaHCO~ 0.2 g CaCI,.2 H,O 0.3 g
649
"O
-r
KC1 KHePO~ NaBr SrCL.6H,O Fe(NH)citrate
0.5 g 0.42 g 0.05 g 0.02 g 0.01 g
Alternatively, the following recipe can be used. NaC1 20.0 g Na~SO~ 3.3 g KC1 0.5 g KBr 0.05 g H~BO~ 0.02 g MgCI~.6HeO 3.0 g Trace minerals (stock solution, in g 1 '): Nitrilotriacetic acid 1.5 g MgSOc7H~O 3.0 g MnSO,.2H~O 0.5 g NaC1 1.0 g FeSO,.7H~O 0.1 g CoCL 0.1 g CaCL.2H~O 0.1 g ZnSO~ 0.1 g CuSO,.5H~O 0.01 g AIK(SO,): 0.01 g H:BO: 0.01 g NaMoO~.2HeO 0.01 g Vitamin stock solution (in g 1 ') (Wolin et al., 1963) Niacin 10 mg Biotin 4.0 mg Pantothenate 10 mg Lipoic acid 10 mg Folic acid 4.0 mg p-Aminobenzoic acid 10 mg Thiamine (B,) 10 mg Riboflavin (B3 10 mg Pyridoxine (B,,) 10 mg Cyanocobalamin (B,:) 10 mg Add 5 ml of the trace element solution and 5 ml of the vitamin mixture to 11 of artificial seawater. Vitamins should be filter-sterilized and added after autoclaving. For the enrichment of chemolithoautotrophs it should not be necessary to add vitamins. Some hyperthermophiles have specific trace metal requirements. For example, growth of Pyrococcus furiosus is stimulated by tungsten (W) (Mukund and Adams, 1991). The salinity within sulfide structures is likely to be higher than that of seawater, and therefore, varying the salt concentration of enrichment media may isolate novel microbes. 650
Electron accepters are usually added at 10 mM and electron donors at concentrations between 0.1 and 0.59~ (w/v). Elemental sulfur is added after autoclaving. Sulfur can be sterilized by steaming for 3 h on three successive days. Na-bicarbonate is also best s u p p l e m e n t e d after autoclaving from a filter-sterilized 5% ( w / v ) stock solution that should be kept under a C O atmosphere. Precipitates may be present after autoclaving m tubes that contain CO, or Na-bicarbonate in their headspace. Shaking can often redissolve those precipitates. Alternatively, the m e d i u m can be autoclaved u n d e r a N, atmosphere and the N~ is exchanged with C O after autoclaving. Organic carbon sources, such as simple sugars, complex carbohydrates, organic acids and alcohols are added at 0.2-0.5~ (w/v). Peptides and proteinaceous substrates are usually added at concentrations between 0.02'.:'i (as a vitamin source) and 0.59; (as a carbon source). Yeast extract or similar substrates at certain concentrations inhibit some thermophiles. Be mindful of this when a m e n d i n g media \xith w~ast extract and try to keep the concentration low.
Special considerations for media for methanogens A general m e d i u m for the enrichn~ent of methanogens and a brief s u m m a r y of the adjustment of the pH by change of the partial pressure ot CO, can be fotmd on the OCM (Oregon Collection of Methanogens) webpage (http://caddis.esr.pdx.edu/OCM/intro.latnal) or see Boone t,t ~11. (1989).
A medium for thermophilic iron reducers The following recipe has been successfully e m p l o y e d for the enrichment of iron-reducing microbes (Kendall and Reysenbach, unpublished). Fe-reducing m e d i u m (g 1 '): NaFtCO, 2.52 ~ NaCI 20 g MgSO~.ZH,O 0.5 g NH;C1 (/.25 g CaCI,-2H~O 0.05 g K,t tPO 4 0.05 g FeSO,.7H:,O 7 mg Na,WO;2H~O (1.3 mg Resazurin Img Fe(lll)citrate 3.67 g Co-enzyme M 0.15 g Dissolve chemicals, adjust p H to 5.8 or desired pH, and dispense 5 ml into Bellco tubes (or serum vials) u n d e r a CO, atmosphere and autoclave. Add 50 1~1 of vitamin solution and 250 t~1 of a 10 mM cysteine solution. After inoculation, the tubes are presstrrized with H, to 20 psi.
651
4,J ¢-
b .r
Preparing anaerobic media See Robb et al. (1995) for detailed anaerobic technique and set up. Some basic approaches are outlined below. Boil medium containing 1.0 mg 1 ' resazurin (redox indicator) and flush with N:, CO, or Ar prior to dispensing into tubes or flasks. Continue flushing with gas when dispensing the medium (Hungate et al., 1969). Alternatively, media can be prepared with oxygen-free water; boiling is then not required. The tubes are sealed with gas impermeable butyl rubber stoppers and crimped with aluminum caps. To remove remaining O~ after autoclaving add sterile Na,S until decoloration of the resazurin indicates a complete reduction of the medium. Alternatively, especially in cases where sulfide should be avoided, the medium can be reduced by adding titanium citrate (Zehnder and Wuhrmann, 1976) until the medium is clear.
Adjustment of pH The pH of the medium has to be adjusted at the incubation temperature adjusting the calibration buffers for higher temperatures. Buffers are added at concentrations between 1-10 mM. Autoclaving can change pH values considerably and pH should be checked and readjusted after autoclaving. The presence of CO~ in the headspace can influence pH. For instructions on pH adjustment in the presence of CO~ see OCM webpage (http://caddis.esr.pdx.edu/OCM/intro.html).
Isolation of thermophiles o n plates Due to the low melting temperature of agar, alternative thermostable solidifying agents are required for growing thermophiles on plates or roll tubes. Both Gelrite (Sigma G1910) and Phytagel (Sigma P8169) have been used for this purpose. The appropriate concentration of the solidifying agent depends on the desired pH (see table below for Gelrite). Gelrite is not stable at a pH lower than 3. For plating highly acidophilic organisms, plates can be prepared with 10% starch (Schleper and Zillig, 1995). pH
Concentration
(g 1 ') 4 5 6 7 8 9
4.92 2.93 1.36 0.64 0.50 0.40
For media preparation, Mg ~" and Ca ~ salts have to be omitted from the medium and are added after autoclaving from a tenfold-filter-sterilized stock solution. If the total concentration of Mg ~' and Ca ~- ions in the
652
m e d i u m is less than 10mM, the concentration has to be adjusted by adding the appropriate a m o u n t of a 1 M MgCI: solution to obtain a final concentration of 10 mM Mg :~. Once the salts are a d d e d the Gelrite will set rapidly and cannot be redissolved. For microbes that are sensitive to high concentrations of Mg ~ ions, as with some Archaea, MgCI~ can be partially replaced with CaCL. Some microbes, such as Sulfiolobus spp, do not grow on surfaces but can be plated in an overlay. For overlays the Gelrite concentration is reduced to 2 g l ', although the appropriate concentration might need to be adjusted empirically. For anaerobes, plates should be poured, streaked and transferred into the incubation jar inside an anaerobic chamber. Even for aerobes, plates should be incubated inside a jar to avoid desiccation due to high incubation temperatures.
Isolation of thermophiles in roll tubes In m a n y cases using roll tubes is preferable to plates, especially for the cultivation of anaerobes in the absence of an anaerobic chamber, or for organisms that require a defined composition of the gases in the headspace. For incubation temperatures up to 70°C agarose (1% w / v ) can be used. For temperatures over 70°C, Gelrite or Phytagel are required. Anaerobically dispense 5 to 10 ml of Gelrite-containing m e d i u m into tubes and stopper, cap and autoclave. Immediately after autoclaving, transfer the tubes to a water bath set at 85-90°C. All solutions that have to be a d d e d after autoclaving (vitamins, phosphates, Mg :* and Ca" salts, inoculum) should be p r e - w a r m e d to the same temperature. Add solutions to one tube at a time, working quickly. The Mg ~' and Ca :' solution must be a d d e d last. Roll the tubes in a tray containing cold water and ice to ensure fast setting of the medium.
Storage of thermophilic isolates and enrichments Many thermophiles can be stored for several months at room temperature or at 4°C u n d e r anaerobic conditions. For long-term storage, cultures should be preserved at -80°C with either 20% glycerol or 15~/~ sucrose as cryoprotectants. Some organisms can be lyophilized and preserved almost indefinitely. We store m a n y of our obligate anaerobes in glass ampules u n d e r gas (see OCM webpage) in liquid nitrogen.
4~ ¢-
b
FUTURE DEVELOPMENTS There will be an increasing need to develop ill situ technologies to measure microbial activity and diversity at deep-sea h y d r o t h e r m a l vents. One area where this is being developed, both for Atlantic and Pacific hydrothermal vents, is through interdisciplinary effort to establish fiber optic cables to provide p o w e r for k)ng-term in situ experin-tents and relay 653
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data back to the surface laboratory. One such proposal is d o c u m e n t e d at http://wwwoneptune.washington.edu. Determining the active populations at deep-sea hydrothermal vents has not been explored extensively. Developing ways to stablize mRNA i~ situ will help address some of these issues. Additionally, basic understanding of microbial dynamics, successional changes and responses to periodic geological and geochemical changes has not been explored in depth. The microbial successional changes that might mirror the invertebrate succession following new eruptive events may help drive some of the methodological d e v e l o p m e n t required to address these basic ecological questions. Furthermore, the rapid d e v e l o p m e n t of microbial genomics and microarray technology for addressing basic ecological questions will influence experimental strategies at deep-sea vents. Finally, efforts that explore innovative methods for microbial sampling at deepsea h y d r o t h e r m a l vents need to be encouraged.
Acknowledgements We thank Melissa Kendall for sharing her iron-reducing recipe. We thank Jeff Seewald and David Butterfield for providing information and photographs of their samplers.
References Atlas, R. M. and Bartha, R. (1993) Microbial Ecology, 3rd edn. The Benjamin/Cummings Publishing Company, Inc. Baross, J. A. and Deming, J. D. (1985). Growth at high temperatures: Isolation and taxonomy, physiology, and ecology. In: Tile Microbiology of Deep-sea Hydrothermal Vents (D. Karl, Ed.), pp. 169-217. CRC Press, Inc., NY. Boone, D. R., Johnson, R. L. and Liu, Y. (1989). Diffusion of the interspecies electron carriers H, and formate in methanogenic ecosystems and its irnplications in the measurement of K,,, for H~ or formate uptake. Appl. Enviros~. Microbiol. 55, 1735-1741. Dempster, E. K., Pryor, K. V., Francis, D., Young, J. E. and Rogers, H. J. (1999). Rapid extraction from ferns for PCR-based analysis. Biotechniques 27, 66-68. German, C. R., Kirk, R. E. and Green, D. R. H. The Buoyant Plume Sampler (BPS): A novel instrument for the investigation of buoyant hydrothermal plumes. (Manuscript in preparation.) Huang, J., Ge, X. and Sun, M. (2000). Modified CTAB protocol using a silica matrix for isolation of plant genomic DNA. Biotechniques 28, 432-434. Hungate, R. E. (1969). A roll tube method for cultivation of strict anaerobes. In: Methods in Microbiology (J. R. Norris and D. W. Gibbons, Eds), pp 117 132. Academic Press, New York. Jannasch, H. W., Johnson, K. S. and Sakamoto, C. M. (1994). Submersible, osmotically pumped analyzers for continuous determination of nitrate in situ. AJtnl. Chem. 66, 3352-3361. Jorgensen, B. B., Zawacki-Leon, X. and Jannasch, H. W. (1990). Thermophilic bacterial sulfate reduction in deep-sea sediments at the Guaymas Basin hydrothermal vent site (Gulf of California). Deep Sea Res. 37, 695 710. Karl, D. M., Taylor, G. T., Novitsky, J. A., Jannasch, H. W., Wirsen, C. O., Pace,
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N. R., Lane, D. J., Olsen, G. S. and Giovannoni, S. J. (1988). A microbiological study of Guaymas Basin high temperature hydrothermal vents. Deep-Sea Res. 35, 777 791. Karl, D. M. (ed) (1995). The Microbiology of DeeF sea Hydrothermal Vents. CRC Press, Inc., New York. Lilley, M. D., Olson, E. J. and Lupton, J. E. (1994). The behavior of CO:, H,, and CH~ in nascent hvdrothermal systems. EOS Trans., Americon Geophysical LIHioJt 75, 618. Lupton, J. E., Delane> J. R., Johnson, H. P. and Tivey, M. K. (1985). Entrainment and vertical transport of deep-ocean water by buoyant hydrothermaI plumes. Nature 316, 621-623. Luther, G. W., Brendel, P. J., Lewis, B. L., Sundb> B., Lefrancois, L., Silverberg, N. and Nuzzio, D. B. (1998). Simultaneous measurement of O,, Mn, Fe, 1-, and S(-II) in marine pore waters with a solid-state voltanrmetric microelectrode. Limmd. OceanoW. 43, 325-333. Marteinsson, V. T., Birrien, J. L., Reysenbach, A. L., Vernet, M., Marie, D., Gambacorta, A., Messner, E, Sleytr, U. B. and Prieur, D. (1999). Thernlococcll5 barophihts sp. nox% a new barophilic and hyperthermophilic archaeon isolated trader high hydrostatic pressure from a deep-sea hydrothermal vent. 1,t. I. Syst. Bocteriol. 49, 351-359. McCollom, T. M. and Shock, E. L. (1997). Geochemical constraints on chemolithautotrophic metabolism by microorganisms in seafloor hydrothermal systems. Ge0chem. Cosmochim. Acta 61, 4375-4391. Mukund S. and Adams, M. W. (1991). The novel tungsten-iron-sulfur protein of the hyperthermophilic archaebacterium, Pyrococclls fllriosus, is an aldehyde ferredoxin oxidoreductase. Evidence for its participation in a unique glycolytic pathway. I. Biol. Chem. 266(22), 14208 16. Reysenbach, A.-L., Banta, A. B., Boone, D. R., Cary S. C. and Luther, G. W. (2000). Microbial essentials at hydrothermal vents.Nature 404, 835. Robb, ET., Place, A.R., Sowers, K.R., Schreier, H.J., DasSarma, S. and Fleischmann, E.M. (Eds) (1995). Archm'a A Laboratory Mamml. Cold Spring Harbor Laboratory Press. Schleper, C., Puehler, G., Holz, I., Gambacorta, A., Janekovic, D., Santarius, U., Klenk, H.-P. and Zillig, W. (1995). Picrophihls gen. nov., faro. no\.:: a novel aerobic, heterotrophic, thermoacidophilic genus and family comprising Archaea capable of growth around pt I O. 1. Bacteriol. 177, 7050-7059. Takai, K., Sugai, A., ltoh, T. and Horikoshi, K (2000). Palaeococcus ferrophilus gen. nov., sp. nov., a barophilic, hyperthermophilic archaeon from a deep-sea hvdrothermal vent chimney l,t. J. S)/st. Evol. Microbiol. 50, 489 500. Taylor, C. D., Wirsen, C. O. and Gaill, E (1999). Rapid microbial production of filamentous sulfur mats at hydrothermal vents. Appl. E,vir. Microbiol. 65, 2253-2255. Von Datum, K. 1.., Edmond, J. M., Grant, B., Measures, C. I., Walden B. and Weiss, R. E (1985a). Chemistry of submarine hydrothermal solutions at 21:~N, East Pacific Rise. Gcochim. Cosmochim. Acta. 49, 2197-2220. Von Damm, K. L., Edmond, J. M., Measures, C. I. and Grant, B. (1985b). Chemistry of submarine hydrothermal solutions at Guavmas Basin, Gulf of California. Gmchim. Cosmochim. Acta. 49, 2221-2237. Wolin, E. G., Wolin, M. J. and Wolfe, R. S. (1963). Formation of methane by bacterial extracts. J. Biol. Chem. 283, 2882-2886. Zehnder, A. J. B. and Wuhrmann, K. (1976). Titanium citrate as a nontoxic oxidation-reduction buffering system for the culture of obligate anaerobes. Science 194, 1165-1166.
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Useful webpages American Type Culture Collection http://www.atcc.org/ Deutsche Stammsammlung f6r Mikroorganismen und Zellkulturen http://www.dsmz.de/ Woods Hole Oceanographic Institution http://www.whoi.edu/ Alvin User manual http: / / www.marine, whoi.edu / ships / alvin / alvin.htm
UNOLS http://www.gso.uri.edu/unols/unols.html Oceanics http://www.generaloceanics.com / index.htm OCM http://caddis.esr.pdx.ed u / Cruise packing lists and other recipes http://caddis.esr.pdx.edu/alr
List of suppliers General Oceanics Inc.
Sigma
1295 N W 16,3 Street Miami FL 33169, USA Tel.: +1 305 621 2882 http://www.businessl .com/genocean/
P.O. Box 14508 St. Louis MO 63178, USA Tel: +1 800 325 3010 ltttp://www.si~lna.sial.com
Deep-sea suppliers for the 'Rosette Syringe Sampler'
Chelex 100 (C7901), PVP 360 (PVP360), Gerite (G1910), Phytagel (P8169)
MJ Research, Inc. 987 Tahoe Boulevard, #106 Incline Village NV 89451, USA TH.: +1 888-735-8437 Fax: +1 617-923-8080 17ttp://www.nor.com/html/instruments /m_m_lab / index.h tml
Mobile molecular laboratory (Model MML 0150)
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Index Note: Page references in italics refer to Figures; those in bold refer to Tables 16S DNA 454 16S rDNA sequence analysis 6()5 16S RNA 427 16S rRNA oligonucleotide probes 200, 208, 217 sequence-based phylogenetic analysis 9 23S rRNA oligonucleotide probes 208, 217 absorptance 523-4 acetate-to-ergosterol (Ac-~ERG) method 359 65 acidophiles 646 acridine orange (AO) 130, 132-4 adenoviruses 564 agar hydrolysis 608 alginate hydrolysis 6(}8 2-alkenyl-4,4-dimethyloxazoline (DMOX) derivatives 601,601,602, 603 ALOt fA, Station 29-32 alpha-complementation 380 Amicon ProFlux M12 ultrafiltration system 55-7 2-amino-2-methylpropanol 601,602 amplified ribosomal DNA restriction analysis (ARDRA) 381,385, 386 template RNA 381-2 anaerobic media 652 antibiotic resistance profiling 555 API 20E test strip 607 API 32A test strip 607 API-ZYM test strip 607 apparent optical properties (AOPs) 523, 528 arachidonic acid (AA) 600, 601 ARB 386, 387 Archaca 9, 14, 15, 17, 376 DGGE of 434 ARDRA analysis 381 2, 385, 386 Ascomycetes 358 aseptic sampling technique 24 astroviruses 564, 565 asymptotic standard error in viability (ASE (V)) 164 ATP 5 as biomarker 253 dissolved (D-ATP) 253-5 intracellular pool turnover 258-6(} Aure~ c~ ccu, aHopha~,~efferens 54 autotrophic bacteria, flow cytometry of 145-6 azoic zone theory 17 Bacillus stearothcrmophilus 629 Bacillus subtilis 111
backscattering ratio 528 bacteria biomass production measurement 227-36 carbon content estimates 137 cell size, determination bv cytometry I45 growth rate measurement 227-36 enumeration 129-52 physiological status of individual cells 175 200 productivity, viral infection and 73 5 sample handling 130-1 sizing 135 6 Bacteria 14, 15, 376 bacterial probe EUB338 223 bacteriophage production, by 'H-thymidine incorporation, viral infection and 68-73 BigDye Terminator Cycle Sequencing Kit 384 binary matrices 454 Bioblanket 644 Biocolumn 644 biologically available phosphorus (BAP) pool 260-4 biomarker profiling 555 biomass production measurement 227-36 BioTools 385 biotraps 644 Bligh and Dver extraction and saponification 602-3 bootstrapping 388-9, 389 bottle effect 27-8, 291 box corers 617 brassicasterol 359 bubble scavenging 20 Buffalo Green Monkey (BGM) cell line 564 Buoyant Plume Sampler 643 burst sizes 75 butterfly baggie sampler 18, 19 C cycle 501 '~C-leucine 469 '*C tracer methodology 25, 32 for primary productivity 4 Q,H: reduction procedure 503-4, 505 6, 507 California Cooperative Ocean fisheries Investigation (CalCOFI) program 23 campesterol 359 carbohydrates, oxidation/fermentation of 609-10 Carlucci, Angelo 7 casein hydrolysis 608 CellQuest 'x' 329 cellular reducing potential 176
657
cyanobacterivory 292 cyanophages 54 PCR in 85 cyclobutyl pyridimidine dimer (CPD) 469 C./:[ ~clastic ts oligot'~pt s 171 cycloheximide 292 CYCLOPS'" 329 Cylitldrothcca 404 Cystofiloba~idimn bisporidii 352 cytopathogenic effects (CPE) 564 CYTOPC 329 C,L/tvlfllo,~a Flavobactcdum 188 Cytvphos,,a Flovvb~wterimu Bacteroides (CFB) 430, 432 Cytophagah's 592, 597 CYTOWin 329, 330 CZCS 526
Centriplus 55, 56 Centriprep 55, 56 cephalexin in DVC 195 Chelex 100 647 Chelo~fia mydtTs 572 chemotaxonomy 605 Chisholm, S.W. 8 chitin hydrolysis 608 Chlorella 63 chlorophyll a measurement 4, 520, 526, 528 remote sensing 533, 534 cholesterol 359 Cholodny-Rossi slides 644 Chrysochromulitla spp. 54 Chytridiomycota 357 CICERO data acquisition system 321 ciproflaxin in DVC 195 Citrobacter 544 cloning / sequencing 9 Clostridimn perfirillgeHs 451,541,542 water quality and548-9 Clostridium spp. 542 CLUSTAL 280 ClustalX 352 cluster analysis 382, 383 Coastal Ocean Observation Laboratory (COOL) 53O Coastal Zone Color Scanner (CZCS) 520 colchicine 292 coliform bacteria indicator species 544 7 colony-forming units (CFUs) 541 Colored Dissolved Organic Matter (CDOM) 534-5 Colwellia 593 Colwellia psycherythreae 597 concentration efficiency 56-7 concentration factor (CF) 56 conductivity temperature-depth (CTD) instrument 21 contact slides 644 Controlled Ecosystem Pollution Experiment (CEPEX) 29 copetrophs 163 Coulter Counter analysis, bacterial sizing by 136 Coulter XL 141 Cmssostrea virj~inica 572 CryptontomTs ozolini 525 Cryptosporidium spp. 572 collection 572-7 IFA system for 577-8 ident{fication 579 PCR-based detection 579-80,581 CTC assav 130, 146-9, 150-1,150, 200 CTC+ celis 146 analysis by flow cytometry 149, 15I cyanine dyes (Cy3, CyS) 179
DAPI (4'6-diamidino-2-phenylindole) 44, 75 bacterial staining bv 140, 179, 183, 184, 187, 200 viral counts by 48 in visualizing bacterial cells 130, 135 DCMU 293 de nov0 pathways 72 deep sea sampling 617-18 see als0 hydrothermal vent microbes degraded DNA, viral 95-6 denaturing gradient gel electrophoresis (DGGE) 9, 284, 425-62 assembly and casting of parallel denaturing gradient gels 448-50 in bacterial spatial and temporal variability 429-30 casting and rtmning of denaturing gradient gels 446 community fingerprinting 217 comparative sequence analysis 460 cycle-sequencing of PCR-products 459-60 DNA amounts to load 452 3 eukaryotic microbial commmfities 434-5 excision of bands and re-amplification 457-9 identification of community members 457 in monitoring population shifts 431-3 pattern analysis 453-61 PCR and RT-PCR for 440-1 principle of separation bv 428 quantification of PCR products 441 running parallel DGGE gels 451 in study of archea, eukaryotes, and viruses 433-4 target sites, sequences and primers 4 4 2 - 3 troubleshooting 450-1 viral communities 435-6 see also PCR-DGGE densitometric analysis 457 depletion method 489-90 DeSoete methods 387
658
Desulfobactcr 429 Desulfobulbtts 429 Desulfovibrio 429, 434 Dewar pyrimidinone 469 diagenetic modeling 489, 490 Dice coefficient 454, 456 differential interference contrast (DIC) microscopy 577 differential metabolic inhibition 8 dilution technique 8 measuring herbivory 293, 294-301,296 dimethyldisulphide (DMDS) derivatization 601,001,603 direct count methods 2 3 direct viable counting (DVC), Kogure method 194-7 dissolved organic material (DOM) 227 distance matrix method 386, 387 diversity indices 457 DMOX derivatives 601,601,602, 603 DNA base composition analysis 605 extraction, yeasts 349 genomic extraction 274-7 microarrays 406 probe hybridizations 281 sequencing 281 DNA:DNA hybridization 605 DNA gyrase inhibition 195 docosahexaenoic acid (DHA) 600, 601 DOM flux 4, 5 downwelling irradiance 522-3 DPAI 306, 307 DTAF 302 ecological approach 615 ecosystem level experiments 28-9 ectoi~ydrolase, bacterial 8 EDTA 140, 647 EGTA 140 eicosapentaenoic acid (EPA) 600, 601 El Nino Southern Oscillation (ENSO) 22 electron microscopy, bacterial sizing by 136 electron transport system (ETS) activity 5 ELISA, DNA damage and 482 Elite'"' 329 energy charge 5 enrichment culture technique 615, 616 enteric protozoan parasites 572-81 enteric viruses 560-71 collection 560-4, 588 detection 564-8, 588 nucleic acid molecular detection 568-71 E~tcrobacter 544 enterococci indicator species 547-8 enteroviruses 555, 560, 564 enzyme assays 8 EPICS 753 321,334, 335
epifluorescence microscopy 6, 131-6 ergosterol 358, 359, 363 ergosterol technique 358, 359, 365 Escherichia coil 110, 111,163, 260, 543, 544 esculin hydrolysis 609 esterase activity 608-9 E~lcarya 14, 18, 376 EurOPA 321 extinction culture isolation 162, 163-71 culture storage 170 1 in diffusion chamber 169-70 location of populations 170 population determination 164 preparation of dilution medium 168-9 properties 171 pure culture production 165 resuscitation 171 viability calculations 164 viability with known number of species 165-8
F~-specific RNA coliphage 549-54 FACSCalibur 318, 332, 334, 335 fastDNAml 387, 388, 389 fatty acid methyl esters (FAME) 600 1,601 filter-concentration method 2 Fisher, Bernhard 2 flow cytometry of autotrophic picoplankton 317-36 advantages and limitations 334-5 analysis 320 calculation 330-2 calibration 332-3 data analysis software 329-30 data storage/archival 333 future directions 335 6 instrumentation 318-19, 319 principles 318 20 protocols 325 9 flow cytometry of bacteria 130, 136 46 basis of method 136 8 counting 140-I protocol 142-6 reference bead standard 141 sample fixation 140 stains 140 Fluoresbrite 141 fluorescein isothiocvanate (FITC) 178, 577 fluorescent in sitH hybridization (FISH) 122, 152, 390, 409 of environmental samples on membrane filters 208-16 fixation / preparation of sediment samples 211-12 fixation of plankton samples 211 fixation for the TSA method 212 hybridization of ceils on rnembrane filters 212-14
659
x qJ
"o
fluorescent in situ hybridization (FISH) (cont.) of environmental samples on membrane filters (cont.) hybridization with HRP labeled probes and TSA 214-15 with rRNA-targeted oligonucleotide probes 207-23 cell brightness quantification by image analysis 221 design and testing 216-22 multi-well glass slides 220 fluorescent microbeads I41 fluorescently labeled bacteria (FLB) measuring herbivory 293, 301-10 disappearance experiments 307-8 uptake experiments 306-7 fluorescently labeled viruses (FLV) 68 tracer dilution using 75-9 fluorescently labeled prey 8 fluorochromes 138, 139 fluorophores 323, 324 FluorX 179 FluoSpheres 141 Forbes, Edward 17 fraction of infected cells (FIC) 74 fraction of mortality due to viral lysis (FMVL) 74 fraction of visibly infected bacterial cells (FVIC) 74 frequence of dividing cells (FDC) method 6-7 fucosterol 359 full cycle rRNA analysis 217 fungal biomass and productivity 357-66 gas chromograph-mass spectrometry (GCMS) technique 600, 602, 604, 605 GCG 280 Gee, Haldane 3 gelatin hydroloysis 607 GelCompare software 382 Genetic Data Environment (GDE) 386, 387, 389 genetically labeled prey 8 GeneTool 385 genomic analysis 9 geometrical radiometry 521-3 Giardia spp. 572 collection 572-7 identification 579 IFA system for 577-8 Glaciecola pmHcea 599 glass plate sampler 20 glutamine synthetase 502 Go-Flo ~ sampling bottles 32, 617 gravity corers 617 grazing, protozoan 8-9 green fluorescent protein (GFP) gene 122 ~H-adenine incorporation method 7
660
~H-labeled compounds 26 3H-leucine 469 incorporation by filter method 228-32 by microcentrifuge method 232-4 ~'H-thymidine 471 incorporation 7 bacteriophage production by 68-73 Hafltia 544 HALE ALOHA 33 tqalophytophthora 366 Hardy continuous plankton record 23 Hawaii Ocean Time-series program (HOT) protocols 29-34, 251 ancillary measurements and experiments 33-4 core parameters 31 discrete depth measurements 30-2 flux and rate measurements 32 high resolution depth profiles 30 light and meterology 33 moored instruments 33 plankton net tows 32-3 site selection 29-30 hepatitis 565 hepatitis A virus 555, 560, 568 Hcterocopsa sp. 525 Heterosigma akashiwo 54 heterotrophic bacteria, flow cytometry of 145-6
heterotrophic potential (V,,,.,,) 4 hexosamine techniques 358 high-pressure pump 624 high-pressure technique, laboratory 616, 617 Hobbie, John 3 Hoescht 33342 140 Hoppe, Hans-Georg 8 hybridization analysis 457 HYDROLIGHT 526, 527 Hydrothermal Fluid Particulate Sampler (HFPS) 641-2, 642 hydrothermal vent microbes 639-54 diversity estimation 647-53 DNA extraction 647-8 elevators vs baskets 640-1 enrichment, culturing, isolation and maintenance 648-53 in situ growth and activity measurement 644-6 in situ sampling devices 641-4 microbial mat and sediment samples 643 rocks, animals and active sulfide structures 643-4 water 641-3 pressure considerations 641 sampling procedures 640-7 sampling vehicles 640 shipboard sample handling 646
in natural bacterial populations 107-8 screening marine bacteria for 111-16 marine prophage induction assay 115-16 natural populations 112 14 prophage induction by viral reduced method 114-15 transduction assay 116 21 in cultured isolates 118 20 in natural populations 121
useful equipment 646-7 hyperpiezopsychrophiles 616 HvPerSort cytometry 336 IFA procedure 577 for CryptosporidiHm sp. 577-8 for Giardia sp. 577-8 imaging cytometrv,, bacterial sizing by 136 immuuomaNHetic separatiou (IMS) 573,576 7 in situ incubation 27 8 iJl situ incubation sampler (ISIS) 27 iH situ PCR 54, 410 itl sitH PCR/RT-PCR/FISH 409-22 ilt vitro PCR/RT-PCR 410 incubation experiments and rate determinations 23-8 Indian Remote Sensing Satellite MOdular Optoelectronic Scanner (MOS) 535 inherent optical properties (lOPs) 523, 524, 528 intergenic spacer region (IGS) 355 IPTG 380 isotope dilution 26, 229 causes 72 isotopic tracers 25-6 Jacard coefficient 456 Johnson-ZoBelI (J-Z) bacteriological sampler 17-18 Jukes Cantor one-parameter model 387 llltlCllS t'OCHlt't'igllllS 364 Kahe, Station 30 kanamycin resistance 117 Kimura two-parameter model 387 kinetic treatment of tracer data 26-7 Kh'bsicllo 544 Koch, Robert 2 Kogure DVC method 194-7 Lambert-Beer's law 526 Laredo sampler 644-6 Lasergene99 352 lecithinase activity 609 leucine incorporation method 7 bacterial measurement by 227-36 Limulus amoebocvte Iysate assay 129 lipase activity 60c} lipopolysaccilaride Limu[us amoebocyte lysate assay of 129 markers 5, 24 LIVE/DEAD BacLight'" bacterial viability kit 191 4 Long-term Ecosystem Observatory (LEO-15) 53O, 533 Lupton sampler 641 lysogen 105 lysogeny advantages 105
M13F primer 385 M13R primer 385 magnesiuna-induced coprecipitation (MAGIC) method 247-51 Mass Selective Detector (MSD) 602 n~aximum likelihood approach 386, 387 8 maximum parsimony 386, 387 membrane integrity 176 Membrex ultrafiltration 112 metabolic inhibitors 293 methanogens, media for 651 microautoradiography 187-91,200 microbial public health indicators 541-55, 543 microbial respiration 5-6 microcentrifuge, }fqetlcine incorporation by 232 4 Micrococcus euryholis 625 Micrococcus lysodeikticlts 451 microelectrodes 644 MICRO-FISH method 188 Micromollas t~ttsillo 54 MIDI system 600 Millipore 55, 56 Mission to Planet Earth (MTPE) program 529 mitomycin C 108, 111, ll2SYBR Gold in lysogen screening 112 Moderate Resolution Imaging Spectroradiometer (MODIS) 529, 535 MOdular Optoelectronic Scanner (MOS) 535 MoFlo cytometry 336 molecular phylogeny 375 90 molecular sequence analysis of yeasts 349-52 12-molybdophosphoric acid (12-MPA) 241 Morisita index 94 most probable number (MPN) assays 55, 57-60 cf plaque assay for enumeration of phycoviruses 62 of psychrophilic bacteria 596 7 Mrakia 355 mRNA analysis, gene expression bv 395-406, 410 applications and performance of assay 405 BOOM method for RNA isolation 401-3 dotting, probing, and quantitating mRNA levels 404-5 GIPS method for RNA isolation 399-401 natural sample collection 398-9 precautions 397-8
661
x
"1:1 era
PAUP' 280, 342, 353, 387, 389 PCR 122, 277-81,410 advantages and disadvantages 582 amplification 9 Cryptosporidium detection 579-80, 581 human viruses and 568 of nitrogenase genes 272-3 of phycoviruses 54 plasmid transduction and 117 primers and controls 273-4 of viral assemblages 85 of yeast 352-4, 355 PCR-DGGE limitations 461-2 reproducibility and sensitivity 460-1 applications 428-9 in spatial and temporal variability of bacterial populations 429-30 in monitoring population shifts 431-3 in study of archea, eukaryotes, and viruses 433-4 eukaryotic microbial communities 434-5 viral communities 435-6 equipment 436 sampling of bacteria 436 extraction of nucleic acids 437 method 437-8 RNA purification 438-9 first strand cDNA 439 PCR and RT-PCR for DGGE 440-1 quantification of PCR products 441 PCR-cycling condifions 444-5 troubleshooting 446 reagent preparation 447-8 Phaeocystis 222 Phaeocystis pouchetii 54 Phaffia 355 phage DSI 118 F116 118 M13 107 P1 107 phagotrophic protists 290 phosphate, radiolabeled (~2po~) 72 phosphorus cycle 239-64, 240, 242-6 phosphorus biologically available (BAP) 260-4 detection in seawater 241-7 radioisotopes ~P 255-6 '~P 255-6 phosphorus-containing compounds detection in seawater 241-7 photoadaptation 300 Photobacteria 3 Phycodnaviridae 54, 435 PCR in 85 phycoviruses, quantification 53-63
MULTIANALYST 441 multidimensional scaling analysis (MDS) 456 multiplicity of infection (MOI), lysogeny and 107, 118 muramic 5 Murphy-Riley method 247, 251 N cycle 501-2 '~Ne procedure 504, 506 7 nalidixic acid in DVC 195 naphthalene dioxygenase 396 Nei and Li coefficient 454 neighbor-joining methods 387, 388 nested PCR 283 nets 16 n!~G (ni~DGK) genes 271 ni~H genes 271,272, 273, 274, 274, 280-1,284, 396 nifHDK genes 271 Niskin baggie sampler 18, 19 Niskin bottles 20-1,617, 643-4 nitrogen fixation 271-84, 503-8 C~H: blockage procedure 509-10 ' N~denitrification 508-12 nitrogenase enzyme 271 nitrogenase gene expression 281 Nitzschia 451 NO~ reductase 502 Nomarski differential interference contrast (DIC) microscopy 577 North Pacific Subtropical Front 22 North Pacific subtropical gyre (NPSG) 23, 29 Northern blotting 281 Norwalk viruses 555, 560 Norwalk-like viruses 568 Nucleic Acid based Sequence amplification (NASBA) 406 nucleic acid markers 24 nucleoid staining technique 183-6, 200 NucliSens ~M401 Ocean Color Monitor (OCM) 535 Ocean Drilling Program (ODP) 640 Ocham's Razor 387 oligobacteria, isolation of 161-72 nomenclature 162-3 oligonucleotide probes 187-91,200 oligonucleotides 385 oligotrophs 163 Oomycota 357, 365 O-ring piston seal 619, 620 Ortho Cytoron Absolute 141 OsmoSampler 643
Palaeococcus ferrophilus 641 particle interceptor traps (PITs) 32 particulate phosphorus 251-5 passive sensors 522
662
PHYLIP 280, 387, 389 phylogenetic analysis 273, 280-1 preliminary steps 385-6 Phytophthora 366 Pi analysis 247-51 uptake/regeneration and DOP production/utilization rates 255-8 PicoGreen 145 picoplankton, autotrophic, flow cytometry of 317-36 piezophilic bacteria, deep-sea 615-34 colony-forming ability assay 629-31 Doryaki method 631 inoculated silica gels formation 630-1 pour tube method 629 silicic acid sol preparation 630 maintenance 631-2 for microbiokGy 618-22 pressure vessels 618-24 sampling 61~18 piezopsychrophilic bacteria 629 PILEUP 280 pipemidic acid in DVC 195 piromidic acid in DVC 195 plankton antiserum binding using immunoprecipitation 475-6 applications 477-80 competitive binding assay (RIA) 477 data analysis 48{1 DNA isol-ation 476-7 immunization schedule 475 immunogen preparation 474 UVB-induced DNA damage 469-82 plaque assays 54, 60 3 cf MPN assay for enumeration of phycoviruses 62 plasmids pQSR50 117 pTYBlue T vector 279 pGEM T vector 279 Polar Front 22 Polaribacter sp. 593, 606 POLarization and Directionality of the Earth's Reflectance (POLDER) 535 PolarotHomzs 593 polioviruses 555 polymerase chain reaction (PCR) see PCR polyphasic taxonomic approach 605 polysaccharides, hydrolysis of 608 polyunsaturated fatty acid (PUFA) 600-1 Pomeroy, L.R. 3 post-collection incubation 24 pour tube mefllod 629 pressure gauges 624 pressure vessels 618-24 culture containers 624 9 direct incubation 629
heat-sealable plastic transfer pipettes 625 plastic bags 625-7, 627 syringes 627-9, 628 test tubes 625 enrichment cultures at high pressure 631 high pressure laboratory instruments 632 sampling with decompression 632 sampling without decompression 632 high-pressure pumps 624 pin-closure 619-22, 619 pressure gauges 624 pressurized temperature gradient (PTG) incubators 633 quick-connect fitting 623 safety issues with high-pressure equipment 633 4 pressurized temperature gradient (PTG) incubators 633 prism dip 20 Prochlorococcus 430 flow cytometry of 321,330, 331-2, 333, 334-5, 336 Prochlorococctts marlines 8, 15 prochlorophytes, discrimination~enumeration bv flow cvtometrv 145 prokary[~tic in s[tu PCR iPI-PCR) 410-22 chemicals, enzymes and supplies 415-16 direct 413 14 gene detection 416-17 iH situ hybridization 419-20 indirect 413, 414, 414 mRNA target detection 418-19 principles 411-15, 412 prokaryotic ilt sit, PCR/FISH (PI-PCR/FISH) 411,414-15 prokaryotic in situ RT-PCR (PI-R%PCR) 411, 413,413, 421 prophage 105 Propidium Iodide (PI) 192 Proteobacteria 221,222 8-Proteobacteria 429 protistan herbiw}ry/bacterivory 289-311 chemical fractionation 292-3 dilution technique 293, 294-301,296 fluorescently labeled bacteria (FLB) 293, 301 1(1 metabolic inhibitors 293 perturbation experiments 291-3 size fractionation 292 Pscttdoalteromotias strain $91 396 pseudolysogeny 105-8, 121 2 Psemhm~omTs spp. 410, 544 Psomhmtolms m'rttxitzosa 109, 118 Psoudomoltas tmtida 411 Psychoflextts torquis 606 psychrophilic bacteria 591-6I 0, 594-5 biochemical tests 607 carbon source and nutritional tests 610
663
x
era
psychrophilic bacteria (cont.) cardinal temperature values 597-600 casein hydrolysis 608 ecophysiological tests 606 enumeration 596-7 fatty acid analysis 600-5 gelatin hydrolysis 607 isolation 592-7 lipolytic enzymes 608-9 oxidation/fermentation of carbohydrates 609-10 phenotypic characterization 605-10 polysacchardies, hydrolysis of 608 storage and cryopreservation 597 pulsed field gel electrophoresis (PFGE) artefactual banding patterns 96, 97 viral fingerprinting by 85-101, 89 pyrimidine(6-4)pyrimidone photoproduct 469-70 Pyrococcus ~uriosus 650
prokaryotic itt situ RT-PCR (P1-RT-PCR) 411, 413, 413, 421 Rhizosoh'Hia 16 Ribosomal Database Project (RDP) 377, 460 Ribosomal Database Project II (RDP) 385 ribosomal RNA (rRNA) cell content 176 RNAzol TM 399 rosette-assisted sampling 21-2 Rosette Syringe Sampler 642, 643 rotating ceramic drum 20 rotaviruses 564, 565 rRNA-targeted oligonucleotide probes with FISH 207-23 RuBisCO large gene 404 R uegeria 432 'S, assays of sulfate reduction rates 489-97 ~SO, methodologies history 490-2 introduction into samples, incubation and termination 492-3 Salmonella 544 Salmonella typhimurium 260, 411 sampler incubation device (SID) 28 sampling platforms 16 satellites, color-sensing 16 scatterance 523-4 screen sampler 20 SeaDAS 529, 531 Sea Wide Field-of-view Sensor (SeaWiFS) 520, 521,535 algorithm 526 SeaWiFS Data Analysis Software (SeaDAS) 529, 531 sediment traps 617 Shannon-Weaver diversity index 457 Shcwaltclla gelidimarina 599 SIMBIOS program 535 sitosterol 359 size-fractionation 8 slurp gun 643 Small Round Structured Viruses (SRSV) 555, 565 Sorensen coefficients 454 Southern hybridization, plasmid transduction and 117 Spartina altcrniflora 358, 364 spin-columns 55 spread plate isolation of oligobacteria 163 square root growth model 597-8, 599 SRP analysis 247 SSU rDNA libraries 377-84 amplification 379 amplified ribosomal DNA restriction analysis (ARDRA) 381,385 ARDRA template RNA 381-2 genomic DNA extraction and isolation 377-8
QIAamp DNAeasy tissue culture procedure 349 Qiagen kit 280 quantitative PCR 54 radioactive isotope tracers 25 radioimmunoassay 469, 473 DNA damage and 482 radio-respiratory method 4 radiotracer method 4 Rayleigh scattering 524 recovery efficiency (concentration efficiency) 56-7 remote sensing, optical 519-36 application 530-5 back image analysis 532-5 obtaining an image 531 processing the image 531-2 biological considerations 525-8 data acquisition 530 future directions 535 hardware requirement 528 9 principle 521-8 reflectance 528 software 529-30 remotely operated vehicles (ROVs) 640-1 Renkonen index 94 reoviruses 564, 565 rescue pathways 72 reverse transcriptase 502 reverse transcription PCR (RT-PCR) 281 3, 410, 542 for DGGE 440-1 human viruses and 569 iI~ situ PCR/RT-PCR/FISH 409-22 in vitro PCR/RT-PCR 410 of nitrogenase genes 272
664
ligation, transformation and screening of clones 380 multitemplate gDNA PCR 379-80 phylogenetic analysis 385-90 putative positive screening 380-1 rarefaction analvsis 383, 384 SSU rDNAs 376 7" sequencing 384-5 stainless steel tray 20 Stathylococcus attreus 163 starch hydrolysis 608 STARFISH protocol 187-8, 191 Sterile Bag Samplers 348 sterile samplers 18-19 stigmasterol 359 Straminipila 357, 365 streptomycin resistance 117 submersible-coupled, in situ sensing and sampling system (SIS3 641 submersibles 640 sulfate reduction rate 490 1 single step chromium reduction in assay 492-7 distillation, trapping and assy of reduced 'S end prod u cts 493-4 '~S assays 489-97 Sulfiolobus 646, 653 SYBR I 40 SYBR-II 140 SYBR Gold, viral counts using 44, 48-9, 79 SYBR Green | 75, 76, 79 enumeration of viruses using 45-9 use in PFGE 99 Sym'chococcus 6, 8, 54, 59, 396, 404 DGGE 435 flow cytometrv 330, 330, 331 lysogeny in 1(i7 ~I.[I1CC]IOCOCCItSUIIICtllIlIS 396 synecology 375-7 SYTO 140 SYTO q 192 SYTO 13 140 bacterial staining 142, 144, 145, 146 Taiwanese Ocean Color hnager (OCI) 535 TAN pool turnover 260 tangential flow filtration 55 TdR method 228, 232 TE buffer 140 temperate phage 105 temperature gradient gel electrophoresis (TGGE) 428 temperature gradient incubator (TGI) 592 terminal restriction fragment length polymorphism (T-RFLP) 9, 284, 389, 461 tetramethyl-rhodamine isothiocyanate (TRITC) 179
Thalassiosira sp. 525 Thermococctts barophihts 641 thennophiles isolation on plates 652-3 isolation in roll tubes 653 storage 653 thermophilic iron reducers, medium for 651 thymidine incorporation 234 5 Tn5 117 topoisomerase 11 inhibition 195 TOPRO 140 TOPRO 1 140 Total Culturable Virus Assav 564 total dissolved phosphate (TDP) measurement bv MAGIC 247, 249-51 tracer dilution, using FLV 75-9 tracer experiments 293-4 transcriptional activity approach 395 transduction, bacteriophage, gene transfer by 108 11 transmission electron microscopy (TEM), viral measurements by 43, 44, 47, 48 of viral particles 67 transmittance 523 4 TREECON 280 trichloroacetic acid (TCA), hot extraction 228-9 Trichodes,zittm I6 Trichodesmium thiebauttii 396 Tri-Reagen t' 399 tritium 26 Triton X 100 140 TrueCount 14l Tyramide Signal Amplification (TSA) 209 tyrosine hydrolysis 609 ultrafiltration, concentration of viruses bv 35-7 ultramicrobacteria 163 u nweighted pair-wise grouping with mathematical averages (UPGMA) 94, 454 6 uricase activity 609 UVB-induced planktonic DNA damage 469-82 principle and methodokGy 472-7 vent cap 644, 04:3 Verity 329 Vibri[~ spp. 544 Vibrio choh'n~c 9 viral productivity (VP) 68-9, 75 viral-like particles (VLP) 110-11 viruses algal, quantification 53-63 fingerprinting by PFGE 85 101, 89 enumeration 43-9 proliferation estimation 67-8{)
665
x ~a "o ¢-
Vital Stain and Probe (VSP) method 177-83 vortex flow filtration 55
yeasts 347-55 collection methods 348 isolation methods 348-9 species identification methods 349-54 strain identification methods 355 YOPRO 140 Yo-Pro I, viral counts by 44, 47-8, 75 YOYO 140
whole ecosystem experimental approach 29 Win MDI 329 X-gal 380 X-ray microanalysis, bacterial sizing by 136
666
