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
O u ° J
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 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
4,a
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om
e~
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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 (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
4~
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I:
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
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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 ( 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
80
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
83
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
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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
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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
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• • • • • •
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*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).
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. m
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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 _,, t,,,..
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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 _ 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 .
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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
010 >,. e-,,~
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
[email protected] lnternet: www.scanalytics.com
Gel analysis software
New England BioLabs
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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
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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
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H©PE -1 (~HSIC Kan + Strep
Treatment
Lysate
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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 (., 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
C QJ 0 .J
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
cO
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).
e, e e e e e
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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2.
3.
4.
5.
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
(pm ~)
(microscopy + C H N analysis)
,m
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S c a n d i n a v i a n coastal waters (X-ray microanalysis)
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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)
0.08+0.02
Oceanic bacterial isolates: {i. 14C acetate uptake)
24.7±7.2
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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~ 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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Molecular Probes, Inc. 4849 Pitchford Avenue Eugene OR 97402 USA 1-541-465-8300
i(~ww.prob~'s.cotH SYTO, SYBR, TOPRO, other blueexcitable fluorochromes, fluorescent bead s Nuclepore Corporation 7035 Commerce Circle Pleasanton CA 94566, USA Membrane filters
Membrane filters Rainin Instrument Co. Inc. Mack Road Box 4026 Woburn MA 01888-4026, USA 1-800-472-4646 E-maih
[email protected] Pipettors
Olympus America Inc. Precision hlstrument Divisioll Two Corporate Center Drive Melville, NY 11747, USA 1-800-445-8236 Epifluorescence microscopes
Sigma Chemical Company P.O. Box 14508 St Louis MO 63178, USA 1-800-325-3010
Other supplies: most standard filtration and microscopy supplies can be obtained from large scientific supply companies such as Fisher Scientific and VWR Scientific Products, which often have company-label products equivalent to, but cheaper than, name-brand products.
Fluorescent beads, CTC
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AO, DAPI Polysciences, Inc. 400 Valley Road Warringtoll PA 18976, USA 1-800-523-2575
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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.
=
-
164
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
1
2 4 10 20 80
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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
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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.
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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
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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.
185
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i~11/ o r~ e0.
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
• • •
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.
186
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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
187
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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:
188
(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• .
189
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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.
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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
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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.
References 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. Braun-Howland, E. B., Danielsen, S. A. and Nierzwicki-Bauer, S. A. (1992). Development of a rapid method for detecting bacterial cells in situ using 16S rRNA-targeted probes. Bioteclmiques 13, 928-934. Bremer, H. and Dennis, P. P. (1987). Modulation of chemical composition and other parameters of the cell by growth rate. In: Escherichia coli and Sahnonella typhinturium: Celhdar and Molecular Biology (E C. Weidhardt, Ed.), pp. 1527-1542, American Society for Microbiology, Washington, DC. Brock, T. D. and Brock, M. L. (1966). Autoradiography as a tool in microbial ecology. Natmv 209, 734-736, Brock, T. D. and Madigan M. T. (1988). Biology q(Microotxanisms, 5th edn. Prentice Hall, Englewood Cliffs, NJ. Carman, K. R. (1990). Radioactive labeling of a natural assemblage of marine sedimentary bacteria and microalgae for trophic studies: an autoradiographic study. Microbial Ecol. 19, 279-290. Choi, J. W., Sherr, E. B. and Sherr, B. E (1996). Relation between presence/absence of a visible nucleoid and metabolic activity in bacterioplankton cells. Limnol. Ocemrogr. 41, 1161-1168. Choi, J. W., Sherr, B. E and Sherr, E. B. (1999). Dead or Alive? A large fraction of ETS-inactive bacterial cells, as assessed by reduction of CTC, can become ETSactive with incubation and substrate addition. Aquatic Microbial Ecol. 18, 105-115. Coallier, J., Prevots, M. and Rompre, A. (1994). The optimization and application of two direct viable count methods for bacteria in distributed drinking water. Can. J. Microbiol. 40, 830-836.
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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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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.
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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
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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
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Southern Micro Instruments Atlanta, GA 30339, LISA; 1-800-241-3312
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Epifluorescence microscopes
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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.
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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.
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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
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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
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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)
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be
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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.)
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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
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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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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/
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High quality filter set for Cy3 Citifluor Ltd 18 Enfield Cloisters Fanshaw Street London N1 6LD, UK Fax: +44 01 227 827724
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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
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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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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 (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
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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,,,~).
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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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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
Extract genomic D N A ]
A m p l i f y nifH by PCR
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Sherr, B. E, Sherr, E. B. and Fallon, R. D. (1987). Use of monodispersed, fluorescently labeled bacteria to estimate in situ protozoan bacterivory. Appl. Environ. Microbiol. 53, 958-965. Sherr, B. E, Sherr, E. B. and Hopkinson, C. S. (1988). Trophic interactions within pelagic microbial communities: indications of feedback regulation of carbon flow. Hydrobiologia 159, 19-26. Sherr, E. B. and Sherr, B. E (1993). Protistan grazing rates via uptake of fluorescently labeled prey. In: Handbook of Methods in Aquatic Microbial Ecology (P. Kemp, B. Sherr, E. Sherr and J. Cole, Eds),pp. 695-701. Lewis Publishers, Boca Raton. Sherr, E. B. and Sherr, B. E (1994). Bacterivory and herbivory: key roles of phagotrophic protists in pelagic food webs. Microbial. Ecol. 28, 223-235. Sherr, E. B. and Sherr, B. E (1997). Phagotrophy in aquatic microbial food webs. In: Manual of Environmental Microbiology. (C. J. Hurst, G. R. Knudsen, M. J. McInerney, L. D. Stetzenbach and M. V. Walter, Eds), pp. 309-316. ASM Press, Washington, D.C. Sherr, E. B., Caron, D. A. and Sherr, B. E (1993). Staining of heterotrophic protists for visualization via epifluorescence microscopy. In: Handbook of Methods in Aquatic Microbial Ecology (P. Kemp, B. Sherr, E. Sherr and J. Cole, Eds), pp. 213-227. Lewis Publishers, Boca Raton. Sieracki, M. E., Haas, L. W., Caron, D. A. and Lessard, E. J. (1987). Effect of fixation on particle retention by microflagellates: underestimation of grazing rates. Mar. Ecol. Prog. Set. 38, 251-258. Simek, K. and Chrzanowski, T. H. (1992). Direct and indirect evidence of sizeselective grazing on pelagic bacteria by freshwater nanoflagellates. Appl. Environ. Microbiol. 58, 3715-3720. Stoecker, D. K. and Capuzzo, J. M. (1990). Predation on protozoa: its importance to zooplankton. J. Plankton Res. 12, 891-908. Stoecker, D. K., Sieracki, M. E., Verity, P. G., Michaels, A. E., Haugen, E., Burkill, P. H. and Edwards, E. S. (1994). Nanoplankton and protozoan microzooplankton during the JGOFS N. Atlantic Bloom Experiment. J. Mar. Biol. Assoc. UK 74, 427-443. Stoecker, R. K., Gustafson, D. E. and Verity, P. G. (1996). Micro- and mesoprotozooplankton at 140°W in the equatorial Pacific: heterotrophs and mixotrophs. Aquatic Microbial. Ecol. 10, 273-282. Taylor, G. T. and Sullivan, C. W. (1984). The use of HC-labeled bacteria as a tracer of ingestion and metabolism of bacterial biomass by microbial grazers. J. Microbiol. Methods 3, 101-124. Taylor, W. D. and Lean, D. R. S. (1981). Radiotracer experiments on phosphorus uptake and release by limnetic microzooplankton. Can. J. Fish. Aquatic Sci. 38, 1316-1321. Tremaine, S. C. and Mills, A. L. (1987). Tests of the critical assumptions of the dilution method for estimating bacterivory by microeucaryotes. Appl. Envilvn. Microbiol. 53, 2914-2921. Weisse, T. and Scheffel-M6ser, U. (1991). Uncoupling the microbial loop: growth and grazing loss rates of bacteria and heterotrophic nanoflagellates in the North Atlantic. Mar. Ecol. Prog. Ser. 71, 195-205. Wikner, J. (1993). Grazing rate of bacterioplankton via turnover of genetically marked minicells. In: Handbook of Methods in Aquatic Microbial Ecology (P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 703-714. Lewis Publishers, Boca Raton. Wikner, J., Andersson, A., Normark, S. and Hagstr6m, ,~. (1986). Use of genetically marked minicells as a probe in measurement of predation on bacteria in aquatic environments. Appl. Environ. Microbiol. 52, 4-8.
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Wright, R. T. and Lebo, M. E. (1987). Dynamics of planktonic bacteria and heterotrophic flagellates in the Parker Estuary, northern Massachussetts. Cont. Shelf Res. 7, 1383-1397. Zobell, C. E. (1943). The effect of solid surfaces upon bacterial activity. ], Bacteriol. 46, 39-56. Zubkov, M. V. and Sleigh, M. A. (1995). Ingestion and assimilation by marine protists fed on bacteria labeled with radioactive thymidine and leucine estimated without separating predator and prey. Micr~bial. Ecol. 30, 157-170.
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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
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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
318
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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.
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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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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
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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
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(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
Fluorescence Absorption color maximum
Emission maximum
Application
Source
(nm)
(nm)
350 358 400 350 347
461 461 420 448 442
DNA; cell cycle DNA; cell cycle Reporter Reporter Reporter
(2) (2) (2) (2) (2)
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
535
617
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)
615 617
DNA, RNA, dead-cell stain Reporter Reporter
670 661 660
Reporter Nucleic acids Reporter
(1) (2) (2)
(2) (2)
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.
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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.
Bead template ~,
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10 0 10 ~ 10 2 10 3 10 4
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RFL
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FALS
SSC
GRFL
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.
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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 (.., (.£,.3
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, 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
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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:
351
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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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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)
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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).
+
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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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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
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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
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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
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100
B
91.7
100
C
43.8
46.7
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27.8
29.4
69.2
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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
°
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 ( 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.
"o c
.x UL .~-
Common equipment and reagents
,oo
•
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
503
4~ °1
Z
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).
505
e" t~
la.'~ C
o.o 4-1 o~
Z
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 (_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.
588
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
t,,t,tt,t
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.
eeeeee
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 ( 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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637
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
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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
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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
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(b)
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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.
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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
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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
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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.
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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)
656
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
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"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
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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
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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
